CHRONIC ETHANOL FEEDING DISRUPTS BOTH LIPID AND GLUCOSE
HOMEOSTASIS IN RAT ADIPOSE TISSUE
by
LI KANG
Submitted in partial fulfillment of requirements
For the degree of Doctor of Philosophy
Thesis Advisor: Dr. Laura E. Nagy
Department of Biochemistry
CASE WESTERN RESERVE UNIVERSITY
May, 2007 CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the dissertation of
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candidate for the Ph.D. degree *.
(signed)______(chair of the committee)
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*We also certify that written approval has been obtained for any proprietary material contained therein. Table of Contents
Title Page i
Committee Sign-off Sheet ii
Table of Contents iii
List of Tables vii
List of Figures viii
List of Abbreviations xi
Acknowledgements xiv
Abstract xvi
Chapter 1 Literature Review 1
1.1 Mechanisms of Ethanol’s Action 2
1.1.1 Enzymatic Metabolism of Ethanol 2
1.1.2 Ethanol’s Action Independent on Enzymatic Metabolism of 4
Ethanol
1.2 Ethanol and Lipid Metabolism 5
1.2.1 Adipose tissue as a regulator of whole-body lipid 5
homeostasis
1.2.2 Regulation of triglyceride metabolism in adipose tissue 6
1.2.3 The effects of ethanol on β-adrenergic receptor-mediated 10
adenylyl cyclase activity and intracellular cAMP
concentration
1.2.4 The effects of ethanol on lipid metabolism 11
iii 1.3 Ethanol and Glucose Metabolism 13
1.3.1 Insulin-mediated glucose transport pathway 13
1.3.2 The effects of ethanol on insulin-mediated glucose 17
metabolism
1.3.3 The effects of ethanol on insulin signaling pathway 19
1.4 Experimental Models of Chronic Ethanol Consumption 21
1.4.1 Lieber-DeCarli liquid diet 21
1.4.2 Tsukamoto-French Rat Model 24
Chapter 2 Research Hypotheses and Objectives 26
2.1 Introduction 26
2.2 Research Hypotheses and Objectives 26
Chapter 3 Dysregulation of Triglyceride Metabolism in Adipose 29
Tissue by Chronic Ethanol
3.1 Introduction 29
3.2 Materials and Methods 30
3.2.1 Materials 30
3.2.2 Animal protocol for determination of triglyceride turnover 31
rates
3.2.3 Measurements of isotope enrichment by gas 32
chromatography-mass spectrometry
3.2.4 Mathematical model for determination of triglyceride 33
turnover rates
3.2.5 Ethanol elimination from blood in rats 34
iv 2 3.2.6 Determination of body fat mass by H2O dilution space 34
3.2.7 β-adrenergic receptor-stimulated in vivo lipolysis 34
3.2.8 Isolation of adipocytes and ex vivo lipolysis assay 35
3.2.9 Hyperinsulinemic-euglycemic clamp and the appearance 35
rate of glycerol (Ra)
3.2.10 Statistical analyses 37
3.3 RESULTS 37
3.3.1 Characteristics of ethanol-fed rats 37
3.3.2 Triglyceride turnover rates 39
3.3.3 β3-adrenergic agonist-stimulated in vivo lipolysis 43
3.3.4 Anti-lipolytic action of insulin ex vivo 46
3.3.5 Anti-lipolytic action of insulin in vivo 48
3.4 Discussion 51
Chapter 4 Disruption of β-Adrenergic Receptor-Stimulated 59
Lipolysis Pathway by Chronic Ethanol
4.1 Introduction 59
4.2 Materials and Methods 60
4.2.1 Materials 60
4.2.2 Animal care and feeding 60
4.2.3 Isolation of adipocytes 61
4.2.4 Ex vivo lipolysis assay and cAMP concentration 62
4.2.5 PDE activity 63
4.2.6 PKA activity 64
v 4.2.7 Western blotting 64
4.2.8 Statistical analyses 66
4.3 Results 66
4.4 Discussion 79
Chapter 5 Disruption of Glucose Disposal by Chronic Ethanol 86
5.1 Introduction 86
5.2 Materials and Methods 87
5.2.1 Materials 87
5.2.2 Animal care and feeding 87
5.2.3 Hyperinsulinemic-euglycemic clamp 88
5.2.4 Glucose utilization rate 89
5.2.5 Radioactivity measurements of plasma and tissue samples 89
5.2.6 Uptake of 2-deoxy-[3H]glucose in isolated adipocytes 90
5.2.7 Statistical analyses 90
5.3 Results 91
5.4 Discussion 96
Chapter 6 Overall Summary and Future Prospects 102
6.1 Overall Summary 102
6.2 Future Prospects 105
Reference List 109
Appendices 125
vi List of Tables
Table 1.1 Macronutrient composition of Liber-DeCarli liquid diet 23
Table 3.1 Rat body weights, plasma ethanol concentration and 38
epididymal fat weights
Table 3.2 Rates of triglyceride synthesis and breakdown 45
Table 3.3 Characteristics of rats in hyperinsulinemic-euglycemic 50
clamp study
vii List of Figures
Chapter 1 Literature Review
Figure 1.1 Hormonal regulation of lipolysis in adipocytes 7
Figure 1.2 Insulin-mediated glucose transport 15
Chapter 2 Research Hypotheses and Objectives
Chapter 3 Dysregulation of Triglyceride Metabolism in
Adipose Tissue by Chronic Ethanol
Figure 3.1 Plasma ethanol decay in rats chronically fed with 40
ethanol for 4 weeks
Figure 3.2 Chronic ethanol feeding decreased lipid content in 41
epididymal adipose tissue
Figure 3.3 Body fat mass in pair- and ethanol-fed rats 42
Figure 3.4 2H-labeling of body water and triglyceride-bound 44
glycerol isolated from epididymal adipose tissue in rats
Figure 3.5 Four-week ethanol feeding decreased β3-adrenergic 47
receptor agonist, CL316,243-stimulated systemic
lipolysis
Figure 3.6 Chronic ethanol feeding impairs the ability of insulin to 49
inhibit lipolysis in isolated adipocytes
Figure 3.7 Plasma glycerol and free fatty acid concentrations 52
during the hyperinsulinemic-euglycemic clamp
Figure 3.8 Chronic ethanol feeding inhibits the anti-lipolytic 53
viii response of adipocytes to insulin in vivo
Chapter 4 Disruption of β-Adrenergic Receptor-Stimulated
Lipolysis Pathway by Chronic Ethanol
Figure 4.1 Chronic ethanol feeding decreased β-adrenergic 67
receptor-stimulated lipolysis in adipocytes isolated
from epididymal fat
Figure 4.2 Chronic ethanol feeding suppressed β-adrenergic 69
receptor-stimulated cAMP accumulation by increasing
cAMP degradation via PDE4
Figure 4.3 Chronic ethanol feeding increased basal activity of 71
PDE4
Figure 4.4 Chronic ethanol feeding did not increase the quantity of 72
immunoreactive PDE4A, PDE4B, or PDE4D isoforms
Figure 4.5 Chronic ethanol feeding decreased β-adrenergic 73
receptor-stimulated PKA activity, but not maximal
PKA activity
Figure 4.6 Phosphorylation of perilipin A and HSL were detected 75
by a phospho-(Ser/Thr) PKA substrate antibody
Figure 4.7 Chronic ethanol feeding reduced β-adrenergic 76
receptor-stimulated phosphorylation of perilipin A and
HSL
Figure 4.8 Chronic ethanol feeding did not affect 78
dibutyryl-cAMP-stimulated phosphorylation of
ix perilipin A or HSL
Figure 4.9 Chronic ethanol feeding impaired signaling 80
downstream of phosphorylation of perilipin A and HSL
Chapter 5 Disruption of Glucose Disposal by Chronic Ethanol
Figure 5.1 Rat blood glucose and plasma insulin levels during 92
hyperinsulinemic-euglycemic clamps
Figure 5.2 Chronic ethanol feeding decreased glucose infusion 93
rate, glucose utilization rate, and percent suppression of
hepatic glucose production during the
hyperinsulinemic-euglycemic clamp
Figure 5.3 Chronic ethanol feeding decreased glucose disposal in 95
adipose tissue, but not in skeletal muscle during the
hyperinsulinemic-euglycemic clamp
Figure 5.4 Chronic ethanol feeding decreased insulin-stimulated 97
glucose uptake in adipocytes isolated from epididymal,
subcutaneous, and omental adipose tissues
Chapter 6 Overall Summary and Future Prospects
Figure 6.1 The current model of chronic ethanol’s effects on lipid 103
and glucose homeostasis
x List of Abbreviations
AC adenylyl cyclase
ACTH adrenocorticotropic hormone
ADA adenosine deaminase
ADH alcohol dehydrogenase
ADRP adipocyte differentiation-related protein
APS adapter protein containing PH and SH2 domain
BSA bovine serum albumin
β-AR β-adrenergic receptor cAMP cyclic AMP db-cAMP dibutyryl-cAMP
CAP Cbl-associated protein
C3G Crk SH3 binding guanine-nucleotide releasing factor
CrkII CT10 regulator of kinase II
CYP2E1 cytochrome P4502E1
[3H]2DG 2-deoxy-D-[1,2-3H]glucose
[3H]2DGP 2-deoxy-D-[1,2-3H]glucose phosphate
DNL de novo lipogenesis
EDTA ethylenediaminetetraacetic acid
EF ethanol-fed
ERK extracellular signal-regulated kinase
FAEE fatty acid ethyl ester
xi FFA free fatty acid
GC-MS gas chromatography-mass spectrometry
GNG gluconeogenesis
GLUT glucose transporter
HEPES N-2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid
HGP hepatic glucose production
HSL hormone-sensitive lipase p-HSL phosphorylated HSL
IL-6 interleukin-6
Ins insulin
IR insulin receptor
IRS-1 insulin receptor substrate-1
Iso isoproterenol
MCP-1 macrophage chemoattractant protein 1
NAD nicotinamide adenine dinucleotide
NADH reduced nicotinamide adenine dinucleotide
N-WASP neutral Wiscott-Aldrich Syndrome protein
PBS phosphate-buffered saline
PDE3B phosphodiesterase 3B
PDE4 phosphodiesterase 4
PDK1 3’-phosphoinositide-dependent kinase 1 p-Peri A phosphorylated perilipin A
xii PF pair-fed
PH pleskstrin homology
PI3K phosphatidylinositol 3-kinase
PIP2 phosphatidylinositol 4,5-bisphosphate
PIP3 phosphatidylinositol 3,4,5-trisphosphate
PKB/Akt protein kinase B aPKC λ/ξ atypical protein kinase C isoforms lambda and zeta
PPARα peroxisome proliferator-activated receptor α
PKA protein kinase A
RBP4 retinol binding protein 4
RIPA radioimmunoprecipitation
R-PIA (-)-N6-(2-Phenylisopropyl)adenosine
SDS sodium dodecyl sulfate
SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis
SREBP-1 sterol regulatory element binding protein-1
TG triglyceride
TGH triacylglycerol hydrolase
TNFα tumor-necrosis factor α
VLDL very low density lipoprotein
xiii Acknowledgements
There are many people who helped to make this thesis possible, for which I would like to express my gratitude.
First and foremost, I would like to gratefully acknowledge the enthusiastic supervision of Dr. Laura E. Nagy throughout my graduate studies. Her inspiration, patience, invaluable guidance, and constant support were essential to my education.
Over the past 4 years, she has trained me in numerous aspects including developing my research abilities, gaining confidence, overcoming difficulties, organizing and writing a scientific publication, and collaborating with other people. And because of her, this graduate study has been a sparkling and unforgettable experience in my life.
I also want to give my deepest thanks to Dr. Steve Previs. As a collaborator and committee member, his contributions, support and kindness are greatly appreciated.
I would also like to thank my other committee members, Dr. Martin Snider, Dr.
Richard Hanson, and Dr. Colleen Croniger for sharing their knowledge and providing helpful suggestions for my thesis project.
A special thanks goes to Becky Sebastian and Jessica Cohen, my lunch buddies, who have provided moral support and comic relief throughout the years. I also express my thanks to Megan McMullen and Brian Pratt for their technical assistance and Dr.
Michele Pritchard and Dr. Abram Stavitsky for sharing their broad research experience, which complemented my project. My sincere gratitude goes to all my colleagues in Dr.
Nagy’s lab; you have provided a stimulating and fun environment in which to learn and grow.
xiv Finally, I have to say “thank-you” to my husband, De, for his special love and endless support. I am forever indebted to my parents, my brother, and my sister who have loved me and cared for me all my life. Thank you all.
This thesis was supported by the NIH grant AA 11876.
xv Chronic Ethanol Feeding Disrupts Both Lipid and Glucose
Homeostasis in Rat Adipose Tissue
Abstract
by
Li Kang
Chronic ethanol consumption induces hepatic steatosis and represents an
independent risk factor for type 2 diabetes, associated with the disruption of lipid and
glucose homeostasis. Adipose tissue acts as a regulator for maintaining whole-body
lipid and glucose homeostasis; therefore, it was hypothesized that lipid and glucose
homeostasis in adipose tissue might be vulnerable to chronic ethanol exposure.
Increases in lipolysis, and the resulting release of free fatty acids into the circulation,
have been associated with the development of insulin resistance and liver injury in other
model systems. Therefore, the effects of chronic ethanol feeding on regulation of triglyceride metabolism in adipose tissue were first investigated. The current study has demonstrated that chronic ethanol feeding to rats increased the turnover rate of triglycerides in epididymal adipose tissue; the rate of triglyceride synthesis was increased
1.9-fold and the rate of triglyceride degradation was increased 3.1-fold over pair-fed controls. Triglyceride degradation is stimulated by β-adrenergic receptor activation and inhibited by insulin. Since chronic ethanol increased the rate of triglyceride degradation, it was first hypothesized that the lipolytic responses of adipocytes to β-adrenergic receptors was increased by chronic ethanol. However, chronic ethanol feeding actually
xvi suppressed β-adrenergic receptor-stimulated lipolysis, both in vivo and ex vivo. This
suppression of β-adrenergic receptor-stimulated lipolysis by chronic ethanol was
associated with inhibited intracellular cAMP accumulation and coincident repression of
cAMP-dependent PKA activation and phosphorylation of perilipin A and HSL. These
data suggest that increased stimulation of lipolysis by β-adrenergic receptors did not
contribute to increased triglyceride degradation after chronic ethanol feeding. In
contrast to the role of β-adrenergic agonist in the activation of lipolysis, insulin is the key
inhibitor of lipolysis. The effect of chronic ethanol on the anti-lipolytic action of insulin
was then studied. Chronic ethanol feeding markedly impaired insulin-mediated
suppression of lipolysis in adipocytes isolated from epididymal adipose tissue, as well as
in conscious rats, during a hyperinsulinemic-euglycemic clamp. Taken together, these
data indicate that chronic ethanol feeding increased the rate of triglyceride degradation
due to a loss in the anti-lipolytic response of adipocytes to insulin.
The effects of chronic ethanol consumption on glucose homeostasis in adipose tissue were also investigated utilizing the hyperinsulinemic-euglycemic clamp technique.
Hyperinsulinemic-euglycemic studies revealed that chronic ethanol feeding to rats decreased whole-body glucose utilization and impaired the ability of insulin to inhibit
hepatic glucose production. Since adipose tissue and skeletal muscle are the two major
organs utilizing glucose in response to insulin, the relative contribution of these two
tissues in mediating impaired glucose utilization after chronic ethanol feeding was further
determined. While glucose disposal in skeletal muscle did not differ between pair- and
ethanol-fed rats, glucose disposal in adipose tissues, including epididymal, subcutaneous,
and omental depots, was decreased in ethanol-fed rats compared to pair-fed rats during
xvii the hyperinsulinemic-euglycemic clamp. These data demonstrate that chronic ethanol
feeding decreased whole-body glucose utilization by impairing the utilization of glucose by adipose tissue, rather than skeletal muscle. In summary, this thesis has demonstrated that chronic ethanol feeding disrupted both lipid and glucose homeostasis in adipose tissue.
xviii Chapter 1
Literature Review
Non-insulin dependent diabetes mellitus (type 2 diabetes) is the most common
metabolic disease prevalent in African Americans, Latinos, North Americans, and Asian
Americans/Pacific Islanders, as well as the aged population (1;2). Type 2 diabetes is
characterized by peripheral insulin resistance followed by an insulin-secretory defect, which results in both postprandial and fasting hyperglycemia, a condition with high blood glucose concentration. Having type 2 diabetes increases the risk for many serious complications, including cardiovascular disease, blindness, nerve damage, and kidney damage (3). Risk factors for type 2 diabetes include family history, hypertension, obesity, sedentary lifestyle, genetics, age, and race (4). Moreover, epidemiological studies suggest that abstinence or heavy consumption of alcohol is also a risk factor for type 2
diabetes. The association between alcohol consumption and type 2 diabetes is complex; studies have demonstrated a U-shaped relationship where moderate drinkers have a
decreased risk for type 2 diabetes, whereas nondrinkers have increased risk, and heavy
drinkers have the greatest risk (5-7). Yet, the exact mechanisms by which chronic heavy
alcohol consumption increases the risk for type 2 diabetes are not well understood; the
disruption of lipid and glucose homeostasis with ethanol exposure likely contribute to the
increased risk (8). Considered as the whole-body lipid storage site, as well as one of the
major organs for glucose transport, white adipose tissue acts as a regulator for
maintaining whole-body lipid and glucose homeostasis. In the current project, we have focused on understanding the effects of chronic heavy ethanol exposure on lipid and
- 1 - glucose homeostasis in white adipose tissue. Here I will review the literature regarding
the mechanisms of ethanol’s action, as well as the pathophysiological effects of ethanol
on lipid and glucose homeostasis in adipose tissue.
1.1. Mechanisms of Ethanol’s Action
Alcohol consumption is causally related with more than 60 different medical
conditions, including hepatic disease, cardiovascular disorders, as well as diabetes
mellitus (9). Although the exact mechanisms by which ethanol consumption is
associated with these pathophysiological conditions are unknown, the consequences of
ethanol’s action can be dependent and/or independent of the enzymatic metabolism of
ethanol in the liver.
1.1.1. Enzymatic Metabolism of Ethanol
Ethanol is primarily metabolized in the liver by three enzymatic pathways. The
enzymatic characteristics, as well as molecular and genetic regulation of these pathways
have been extensively reviewed (10-13). Alcohol dehydrogenase (ADH), the major
enzyme in the liver oxidizes alcohol to form acetaldehyde, with the reduction of
nicotinamide adenine dinucleotide (NAD) to its reduced form (NADH). ADH is
predominantly expressed in the liver, but other tissues, including gastric mucosa and
brown adipose tissue also express ADH (11;14). Oxidation of ethanol by ADH accounts
for the bulk of ethanol metabolism at low concentrations of blood ethanol due to the low
Km of this enzyme (0.2-2 mM). Because of the generation of NADH and the subsequent
changes in NAD+/NADH ratio, oxidation of ethanol by ADH leads to a shift of redox
- 2 - state in hepatocytes, which further impacts on intermediary metabolism in the liver. For
example, the increased NADH depresses the activity of the citric acid cycle, a cycle that
normally provides NADH for the respiratory chain through oxidation of two carbon
fragments derived from fatty acids (15). However, as the organism adapts to continued
exposure to ethanol, the shift of redox state tends to be normalized (16); therefore, the
functional consequences of ethanol exposure result from more than just changes in the
redox state of the hepatocytes (11).
Acetaldehyde, the major metabolite of ethanol metabolism by ADH is toxic to the
function of hepatic mitochondria (17). Increased concentration of acetaldehyde
significantly reduces the oxidative capacity of liver mitochondria including the oxidation of fatty acids and acetaldehyde (18;19). Moreover, incubation of hepatocytes with acetaldehyde depresses the transport of fatty acids into the mitochondria by decreasing the activity of palmitoyltransferase I (20). Therefore, not only the increased
NADH/NAD+ ratio but also acetaldehyde, the two products of ethanol metabolism by
ADH, inhibits fatty acid oxidation.
The second major pathway for ethanol elimination is the microsomal ethanol
oxidizing system catalyzed by cytochrome P4502E1 (CYP2E1). Activity of CYP2E1 in
the liver is induced in chronic alcoholics, and this pathway is known responsible for the
detoxification of excessive amounts of ethanol consumption due to a high Km of CYP2E1
(10-15 mM) (11). In addition to ethanol metabolism, another important feature of
CYP2E1 is its extraordinary capacity to convert many xenobiotics to highly toxic metabolites. These agents include industrial solvents, anesthetic agents, and commonly used medications, which explains the increased vulnerability of alcoholics to these
- 3 - substances (11). Moreover, metabolism of ethanol via CYP2E1 also results in the
production of several reactive oxygen species, including superoxide, hydroxyl radical,
and other free radicals, which are thought to contribute to ethanol-induced liver injury
(11). As with ADH, CYP2E1 is also primarily expressed in hepatocytes (11); however,
additional sites of CYP2E1 expression include Kupffer cells in the liver (21) and white
adipose tissue (22).
The final pathway for ethanol metabolism is a nonoxidative pathway catalyzed by
fatty acid ethyl ester (FAEE) synthase, leading to the formation of fatty acid ethyl esters
(23). FAEEs are esterification products of ethanol and fatty acids; there have been many
reports suggesting that FAEEs are cytotoxic (23). The organs most frequently damaged
by ethanol abuse, the pancreas and liver, have the highest concentrations of both FAEE
and FAEE synthase (24). FAEE is also currently developed as a marker for both acute
and chronic ethanol intake (23).
1.1.2. Ethanol’s Action Independent on Enzymatic Metabolism of Ethanol
Ethanol not only produces its effects via enzymatic metabolism and its metabolites,
but also modulates cellular events by directly acting on membrane proteins and lipids. In
the brain, ethanol inhibits the ATP-gated ion channel by interacting with a small
hydrophobic pocket on the membrane receptors of central nervous system neurons (25).
In addition to specific interactions with membrane proteins, it is also hypothesized that
ethanol may interact directly with membrane lipids (26). Several mechanisms of ethanol’s action on membrane lipids have been proposed, including membrane volume occupation (27) and expansion (28), altered membrane fluidity (29), and lipid phase
- 4 - transitions (30). It should be noted, however, the functional effects of ethanol ultimately result from alterations in protein function regardless interacting with membrane lipids or proteins (26).
Ethanol also disrupts cell function by a specific mechanism of the formation of
phosphatidylethanol by phospholipase D, an enzyme that normally catalyzes the
hydrolysis of phospholipids to phosphatidic acids (31). Phosphatidylethanol is formed at the expense of the normal lipid product, phosphatidic acid and phospholipase D-mediated signal transductions in the presence of ethanol. Significant amount of phosphatidylethanol have been found in rat brain, liver, and other tissues after chronic ethanol consumption (32), which accumulates in cell membranes, potentially affecting membrane-associated processes.
1.2. Ethanol and Lipid Metabolism
1.2.1. Adipose tissue as a regulator of whole-body lipid homeostasis
Adipose tissue is a specialized connective tissue that functions as the major storage
site for fat in form of triglycerides. Serving as an energy reserve, adipose tissue
synthesizes triglycerides when energy intake exceeds energy output. During fasting or in
response to infection and inflammation, adipose tissue mobilizes free fatty acids and
glycerol providing other tissues with metabolites and energy substrates (33). Free fatty
acids released from adipose tissue are transported into other tissues, where they are either
oxidized to generate energy or stored as organ fat in form of triglycerides or cholesterol.
In addition, free fatty acids also act as endogenous ligands for a number of transcription factors in the nuclear receptor family, such as peroxisome proliferator-activated receptor
- 5 - α (PPARα) (34). Fatty acid-bound PPARα heterodimerizes with retinoid-X receptor and
binds to a peroxisome proliferator response element in the promoter of the target genes
involved in mitochondrial and peroxisome β-oxidation (34).
Glycerol release from adipose tissue via lipolysis provides metabolic substrates for other tissues. In the liver, glycerol is phosphorylated by glycerol kinase yielding
glycerol-3-phosphate. Glycerol-3-phosphate is further oxidized to dihydroxyacetone
phosphate, either providing energy through glycolytic pathway or acting as the substrates
of synthesis of lipid (lipogenesis) and synthesis of glucose (gluconeogenesis).
1.2.2. Regulation of triglyceride metabolism in adipose tissue
Triglyceride metabolism in adipose tissue is tightly regulated by a number of
hormones. It has been well established that lipogenesis in adipose tissue is stimulated by
insulin through upregulation of lipogenic enzymes including acetyl-CoA carboxylase and
fatty acid synthase (35;36). Recent studies show that the stimulation of lipogenic
enzymes by insulin is associated with the activation of sterol regulatory element binding
protein-1 (SREBP-1), the transcription factors that bind to an E-box motif in the promoter
of fatty acid synthase gene (37).
Similarly, lipolysis in adipocytes is also regulated by hormones, such as ACTH
(adrenocorticotropic hormone), epinephrine, norepinephrine, and insulin (38) (Fig 1.1).
Catecholamine-induced lipolysis is well characterized, initiated by stimulation of β-
adrenergic receptors, which are coupled to activation of adenylyl cyclase by the
heterotrimeric Gαs protein, which in turn converts ATP to cyclic AMP (cAMP). cAMP- dependent protein kinase A (PKA) then phosphorylates two main targets, hormone-
- 6 - β-AR IR
PDK PIP3 PIP2 AC IRS-1 Gs PI3K PKB/Akt PDE3B
PDE4 ATP HSL AMP cAMP
PKA P L P HS PE RIL IP P IN P A TR A IN I P IP E GL IL ID Y ER R Li CE P CE pi RI LY d DE IG dr TR op FFA let GLYCEROL
Figure 1.1 Hormonal regulation of lipolysis in adipocytes.
Lipolysis in adipocytes is stimulated by activation of β-adrenergic receptors and inhibited by activation of insulin receptors. The convergence of these two pathways is to regulate the intracellular concentration of cAMP, which activates protein kinase A and subsequently stimulates the release of glycerol and free fatty acids by phosphorylation and activation of hormone sensitive lipase and perilipin. β-AR, β-adrenergic receptor;
AC, adenylyl cyclase; cAMP, cyclic AMP; PKA, protein kinase A; HSL, hormone sensitive lipase; FFA, free fatty acid; IR, insulin receptor; IRS-1, insulin receptor substrate-1; PI3K, phosphatidylinositol 3-kinase; PIP2, PI 4,5-bisphosphate; PIP3, PI
3,4,5-trisphosphate; PDK, 3’-phosphoinositide-dependent kinase 1; PKB/Akt, protein kinase B; PDE3B, phosphodiesterase 3B; PDE4, phosphodiesterase 4.
- 7 - sensitive lipase (HSL), the primary lipase responsible for hydrolysis of triglycerides, as
well as perilipin A, the coating protein of lipid droplets. In unstimulated adipocytes,
perilipin A functions as a barrier to lipolysis because of its location on the surface of lipid
droplets, preventing the interaction of HSL with the lipid droplet (39). In response to β-
adrenergic activation, phosphorylated perilipin A undergoes a conformational change,
which is essential for proper translocation of HSL from the cytosol to the surface of lipid droplets and subsequent attachment to triglycerides, leading to initiation of triglyceride hydrolysis (39;40).
Phosphorylation of HSL by PKA occurs at three sites, the serines 563, 659 and 660,
both in vitro and in primary rat adipocytes. Phosphorylation of Ser-659 and -660 is required for in vitro activation as well as translocation of HSL from the cytosol to the lipid droplets, whereas the role of the third PKA site remains elusive (38). In addition to the three PKA phosphorylation sites, one additional phosphorylation site in HSL, Ser-600 has been proposed to be phosphorylated by extracellular signal-regulated kinase (ERK)
1/2 (41). Evidence suggests that catecholamines not only can activate PKA, but also
ERK1/2, the mitogen-activated protein kinase. It has been shown that this pathway is
activated through β3-adrenergic receptor coupling to Gi, raising the possibility that dual
Gs/Gi-protein coupling to this receptor allows PKA and ERK1/2 to work in concert to
stimulate lipolysis (42).
Although HSL is considered as the principle lipase of lipolysis in adipose tissue, studies with HSL-deficient mice reveals the presence of one or more additional intracellular triglyceride lipase (43). Studies of lipolysis performed in isolated adipocytes show that catecholamine-stimulated glycerol release is almost completely blunted in
- 8 - HSL-deficient adipocytes compared with wild-type cells (43;44). Basal lipolysis,
however, is unaffected in HSL-deficient mice (43). Studies using a functional proteomics
approach to detect non-HSL lipase(s) in mouse white adipose tissue reveal that
carboxylesterase 3 (triacylglycerol hydrolase (TGH)) is also a principal lipase of white adipose tissue (45). TGH is responsible for a major part of non-HSL lipase activity in
white adipose tissue in vitro and may mediate some or all HSL-independent lipolysis in
adipocytes (45).
In contrast to the stimulatory response of catecholamines, insulin is the key counter
regulatory hormone against β-adrenegic receptor-mediated activation of lipolysis (38).
Insulin binds to insulin receptor, leading to the autophosphorylation of the receptor and
phosphorylation of insulin receptor substrate-1 (IRS-1). Phosphorylated IRS-1 activates
PI 3-kinase by binding to the regulatory p85 subunit and releasing the catalytic p110
subunit. The catalytic p110 subunit of PI 3-kinase then catalyzes phosphorylation of
plasma membrane-associated PI 4,5-bisphosphate (PIP2) to produce PI 3,4,5-
trisphosphate (PIP3) in the cellular membrane. The formation of PIP3 in the plasma
membrane generates recognition sites for a number of proteins containing pleskstrin
homology (PH) domain including the serine/threonine kinases 3’-phosphoinositide-
dependent kinase 1 (PDK1) and protein kinase B (PKB/Akt). Activated PKB/Akt further
phosphorylates membrane-associated phosphodiesterase 3B (PDE3B) on serine 302,
resulting in decreased cAMP production and inhibition of catecholamine-induced
lipolysis (46).
- 9 - In contrast to activation of PDE3B by insulin, phosphodieasterase 4 (PDE4), another isoform of PDEs that degrade cAMP to AMP, is regulated by PKA- and mitogen- activated protein kinase-mediated phosphorylation (47).
1.2.3. The effects of ethanol on β-adrenergic receptor-mediated adenylyl cyclase activity
and intracellular cAMP concentration
Activation of adenylyl cyclase by hormones and neurotransmitters with the
subsequent generation of cAMP mediates a number of intracellular processes. Ethanol
has a profound impact on β-adrenergic receptor-mediated adenylyl cyclase activity and
intracellular cAMP concentration in varying cell types (48). The in vitro addition of
pharmacologically relevant concentration of ethanol stimulates adenylyl cyclase activity
in striatal (49) and cerebral cortical tissues (50), but decreases the fold stimulation of
adenylyl cyclase activity by isoproterenol, an agonist of β-adrenergic receptor, in
membranes prepared from lymphoma cells (51). Chronic administration of ethanol,
however, causes a desensitization of β-adrenergic receptor-cAMP system. In rodent
brains, chronic ethanol exposure reduces the stimulation of cAMP concentration by
norepinephrine (52). Similarly, isoproterenol-stimulation of adenylyl cyclase activity is
also decreased in cerebral cortex after chronic ethanol administration (53).
β-adrenergic receptor-cAMP system is not the only target of desensitization by
chronic administration of ethanol. Long-term exposure of NG108-15 neuroblastoma-
glioma cells to ethanol results in heterologous desensitization of adenosine receptor- and
prostaglandin E1 receptor-dependent cAMP signal transductions (54).
- 10 - It has been proposed that the chronic ethanol-induced desensitization of receptors is associated with a decrease in the expression of heterotrimetric Gαs protein and its functional activity in many cell types, such as neural cells (55), murine lymphoma cells
(56), and developing rat hippocampus (57). However, other studies investigating different cell types, including rat skeletal muscle cells (58) and rat adipocytes (59), show an increased expression of Gαs protein, coupled with increased intracellular cAMP concentrations. Taken together, these data are indicative of different cell specific effects of chronic ethanol on receptor-dependent adenylyl cyclase-cAMP signal transduction.
1.2.4. The effects of ethanol on lipid metabolism
Accumulation of fat in the liver is the most common disturbance of lipid metabolism produced by ethanol. Hepatic steatosis or alcoholic fatty liver is characterized as the earliest stage of ethanol-induced liver injury, which is followed by the development of inflammation, hepatocyte necrosis/apoptosis, fibrosis and finally cirrhosis. Ethanol- induced fatty liver can be detected both morphologically and biochemically by the increased lipid content of liver. Under controlled conditions, a 25-fold rise of triglyceride concentration is observed in men when ethanol is given as a supplement to normal diet
(60). Ethanol consumption may cause malnutrition because ethanol, as a substantial source of energy, often displaces normal nutrients, such as folate, thiamine, and other vitamins (11). Ethanol-induced malnutrition was first postulated as the mechanism of alcoholic liver disease. However, rats fed with a nutritionally adequate liquid diet supplemented with ethanol develop an obvious fatty liver, suggesting that the pathogenesis of ethanol-induced fatty liver involves not only malnutrition but also a
- 11 - direct effect of ethanol on the liver, most likely due to the effect of ethanol on intermediary metabolism in the liver (61).
One of the important metabolic disorders produced by ethanol metabolism is a
disruption of lipid homeostasis in the liver. ADH-mediated ethanol metabolism causes
redox changes in the liver which promote steatosis by stimulating the synthesis of fatty
acids and opposing their oxidation (61). Moreover, increased fatty acid uptake into the
liver (62), increased esterification of fatty acids in the liver (63), increased synthesis of
cholesterol in the liver (64), and decreased secretion of liver lipid in the form of very low
density lipoprotein (65) are also thought to contribute to the pathogenesis of ethanol-
induced fatty liver.
Alcoholic fatty liver is often associated with hyperlipidemia. Studies both in human
and in animals indicate that ethanol exposure leads to the development of hyperlipidemia characterized by elevated plasma triglyceride, cholesterol, and free fatty acid
concentrations (60). After an acute administration of a single dose of ethanol, healthy
subjects exhibit increased triglycerides in the plasma (66). Prolonged exposure to ethanol
exaggerates the effects of ethanol on plasma triglycerides. Daily consumption of ethanol for 3-5 weeks significantly increased plasma triglyceride concentrations in both human
and animal models (67;68). The ethanol-induced hyperlipidemia in humans is usually transient; serum lipid concentrations return to normal levels after 2-3 weeks of ethanol
consumption in alcoholic volunteers despite continuation of or even an increase in the alcohol intake (60). Conversely, some individuals under particular conditions, such as obesity, exhibit marked and sustained alcoholic hyperlipidemia (69). In addition,
- 12 - extended administration of ethanol in rodents up to 6 weeks still show elevated
concentrations of triglycerides, cholesterol, and free fatty acids in the circulation (70).
Although chronic ethanol feeding disrupts lipid homeostasis in the liver, as well as at
the whole body level, and adipose tissue acts as a regulator of whole-body lipid
homeostasis, the effects of chronic ethanol on the regulation of lipid metabolism in
adipose tissue has been barely investigated. While the in vivo effects of chronic ethanol
on triglyceride metabolism in adipose tissue is unknown, studies of de novo lipogenesis
with labeled glucose in dissected adipose tissue indicate that chronic ethanol feeding
increases the rate of de novo lipogenesis (71). In addition, in adipocytes isolated from epididymal adipose tissue in rats chronically fed with ethanol, both basal and β-
adrenergic receptor-stimulated lipolysis are decreased compared to those in adipocytes
isolated from their counterpart controls; however, the anti-lipolytic effect of insulin and
the adenosine pathway are unchanged by chronic ethanol feeding (72).
1.3. Ethanol and Glucose Metabolism
1.3.1. Insulin-mediated glucose transport pathway
Glucose transport is a highly regulated process that allows for the control of blood
glucose level and normal cell growth and maintenance. A family of specialized
transporter proteins facilitates the transport of glucose across plasma membranes in an
energy-independent manner (73). Six mammalian glucose transporter isoforms (GLUT1-
5 and GLUT7) have been identified, as reviewed regarding their tissue or cell-specific
expression, distinct efficiencies and kinetics for transporting glucose, as well as
differential regulation of the GLUT protein expression (73).
- 13 - Of the 6 isoforms of GLUT proteins, GLUT4 is the isoform expressed only in
adipocytes and muscle cells, the two major sites of glucose transport in response to
insulin (74;75). As with other glucose transporters, GLUT4 belongs to the 12 transmembrane segment transport superfamily (73). Although the three-dimensional
structure of GLUT4 is unknown, it has been proposed that at least five of the
transmembrane helices cluster together to form an aqueous pore, which transports
glucose via a conformational change from an outward to an inward facing conformation
(73;76).
Although adipose tissue and skeletal muscle express other glucose transporter
isoforms, most notably GLUT1, it has been demonstrated that GLUT4 is responsible for insulin-stimulated glucose transport (73). Insulin increases glucose transport in fat and muscle cells by stimulating the translocation of GLUT4 from intracellular sites to the plasma membrane (77). GLUT4 is found in vesicles that continuously cycle from intracellular storage compartments to the plasma membrane. Activation of insulin receptor by insulin binding triggers a dynamic redistribution of GLUT4 protein via increasing the rate of GLUT4 vesicle exocytosis, and via slightly decreasing the rate of internalization (78). Trafficking of GLUT4 vesicle to the cell surface is followed by tethering, docking, and fusion of the vesicles with the plasma membrane, allowing the extracellular exposure of GLUT4 protein and subsequent glucose transport (79).
Insulin stimulates GLUT4 translocation through two independent signaling pathways,
the IRS-PI 3-kinase signaling pathway (Signal 1), and the CAP-Cbl signal pathway
(Signal 2) (79) (Fig 1.2). The IRS-PI 3-kinase pathway is well established and extensively reviewed (79). Phosphorylation of insulin receptor and IRS by insulin
- 14 - Glucose
IR Lipid rafts IR
PDK PIP3 PIP2 Flotillin APS CAP P Cbl PI3K CrkII P IRS C3G
PKB/Akt aPKCλ/ξ TC10 AS160
Signal 1 Signal 2 GLUT4 Storage Compartments
Figure 1.2 Insulin-mediated glucose transport.
The insulin-dependent GLUT4 translocation requires two independent signal pathways; the IRS-PI3-kinase signaling pathway (Signal 1), and the CAP-Cbl signal pathway
(Signal 2). IR, insulin receptor; IRS, insulin receptor substrate; PI3K, phosphatidylinositol 3-kinase; PIP2, PI 4,5-bisphosphate; PIP3, PI 3,4,5-trisphosphate;
PDK, 3’-phosphoinositide-dependent kinase 1; PKB/Akt, protein kinase B; CAP, Cbl- associated protein; APS, adapter protein containing PH and SH2 domain; CrkII, CT10 regulator of kinase II; C3G, a guanine nucleotide exchange factor; aPKC, atypical protein kinase C. Adapted from Kanzaki M. (79).
- 15 - binding activates PI 3-kinase. The catalytic subunit of PI 3-kinase then catalyzes the
conversion of PIP2 to PIP3 in the cellular membrane, which in turn stimulates the activation of PDK1 and its downstream activation of PKB/Akt. Activated Akt/PKB further phosphorylates AS160, a Rab GTPase-activating protein, whose phosphorylation has been shown to be required for insulin-stimulated GLUT4 translocation (80). In addition to PKB/Akt, activation of the atypical protein kinase C isoforms lambda and zeta
(aPKC λ/ξ) is also involved in the process of insulin-stimulated GLUT4 translocation
(81;82).
Although the PI 3-kinase pathway is required for insulin-stimulated glucose uptake,
recent data suggest that it is not sufficient to stimulate glucose transport (83). Baumann
et al. and Chiang et al. have identified a second signaling pathway involved in insulin- stimulated glucose transport. In this pathway, GLUT4 translocation requires activations of the proto-oncogene Cbl and the small GTPase TC10 (83;84). Contrasting with the
IRS-PI 3-kinase pathway, autophosphorylation of insulin receptor results in the binding of the adapter protein, APS (adaptor protein containing a PH and SH2 domain). The association of APS with the insulin receptor then leads to the phosphorylation and association of Cbl, due mainly to the associated protein CAP. CAP binds to Cbl via its three SH3 domains in its carboxyl terminus, as well as flotillin, a lipid raft resident protein via its Sorbin Homology domain in amino terminus. The interaction between
CAP and flotillin results in the recruitment of CAP/Cbl protein complex into lipid rafts.
Phosphorylated Cbl then recruits a signaling complex containing the adaptor protein
CrkII and the guanine nucleotide exchange factor C3G into the lipid rafts microdomains.
C3G stimulates the exchange of GTP for GDP on small G-proteins including TC10, a
- 16 - Rho family small G-proteins present in the lipid rafts. The spatially compartmentalized
TC10 generates signals that contribute to insulin-induced glucose transport. These signals include the activation of exocyst complex that regulates tethering/docking of
GLUT4-containing vesicles, activation of atypical PKC λ/ξ which is also a downstream target for IRS-PI 3-kinase pathway, as well as activation of neutral Wiscott-Aldrich
Syndrome protein (N-WASP) and actin polymerization which has been shown to be required for GLUT4 translocation (79).
However, recent studies have questioned the role of CAP-Cbl signaling pathway in the regulation of GLUT4 translocation and insulin-mediated glucose transport. In APS- deficient mice or 3T3-L1 adipocytes, neither insulin-stimulated glucose uptake nor
GLUT4 translocation was impaired compared to APS-sufficient mice or cells (85).
Furthermore, selective depletion of CAP, Cbl isoforms, or CrkII in 3T3-L1 adipocytes using small interfering (si) RNAs failed to attenuate insulin-stimulated glucose transport or GLUT4 translocation, suggesting that CAP-Cbl pathway may be not required for insulin signaling to GLUT4 transporters (86).
1.3.2. The effects of ethanol on insulin-mediated glucose metabolism
Epidemiologic studies from numerous disparate populations reveal that ethanol consumption is associated the incidence of type 2 diabetes in a U-shaped fashion.
Individuals with daily moderate intake of alcohol show a decreased risk for type 2 diabetes, whereas abstainers have increased risk, and heavy drinkers have the greatest risk (5-7). Research is in progress to explain this observation in molecular and nutritional
- 17 - aspects. Heavy ethanol intake has been shown to disrupt glucose homeostasis, which
likely represents a specific mechanism of increased risk for type 2 diabetes.
Conditions associated with heavy alcohol consumption include both hypoglycemia,
where blood glucose concentration is low because of impaired hepatic gluconeogenesis
and glucogenolysis, and hyperglycemia, where blood glucose concentration is high
because of insulin resistance (87). Ethanol-associated hypoglycemia occurs in subjects
on a low-carbohydrate diet or in fasting subjects who miss a meal or two while drinking
(88). After the consumption of alcohol for 48 hours, hepatic gluconeogenesis is
decreased by 45% in overnight-fasted humans (89). While normal blood glucose can not
be maintained by hepatic gluconeogenesis, glycogen stored in the liver starts releasing
glucose via glycogenolysis. However, glycogenolysis is also impaired by ethanol
consumption (89). When glycogen in the liver is depleted, hypoglycemia occurs. Yet,
even when ethanol consumption is combined with a meal, reactive hypoglycemia occurs
in 2 to 3 h after the meal (90). Ethanol intake with a carbohydrate-rich meal initially
increases blood glucose level since ethanol is the preferred fuel at the expense of glucose utilization. In order to correct the initial hyperglycemia, excessive amounts of insulin or hyperinsulinemia is developed, which drives the blood glucose down, further causing hypoglycemia.
Ethanol-associated hyperglycemia is associated with impaired insulin-stimulated
glucose utilization. With one moderate load of ethanol, also defined as acute intake of
ethanol (0.6-1 g/kg) in healthy men, hyperinsulinemic-euglycemic clamp studies show
that ethanol decreases glucose oxidation and decreases whole-body glucose utilization
(91-93). However, after an extended period of ethanol consumption, defined as chronic
- 18 - moderate use of alcohol, hyperinsulinemic-euglycemic clamp studies shows enhanced
insulin-stimulated glucose uptake compared with non-drinkers (94;95). Studies including
chronic heavy drinkers or alcoholics, however, indicate decreased insulin sensitivity
during a hyperinsulinemic-euglycemic clamp (96). Interestingly, experiments carried out
with animals reveal the similar results. Chronic heavy ethanol exposure to rats decreases
whole-body glucose utilization during the hyperinsulinemic-euglycemic clamp (58;97).
Since adipose tissue and skeletal muscle are the two major sites utilizing glucose as the
energy fuel in response to insulin, the effects of ethanol on the glucose metabolism in
these particular tissues have been studied. Chronic ethanol exposure to rats decreases
insulin-stimulated glucose uptake in isolated adipocytes (59;98). However, the effects of chronic ethanol on insulin-stimulated glucose transport in muscle are controversial.
While Wan et al. reported that chronic ethanol feeding of rats at 5g/kg and 2.5g/kg for 20
weeks significantly decreases insulin-stimulated glucose uptake in isolated skeletal
muscle (58), Wilkes et al. reported that rats fed with Lieber-DeCarli ethanol liquid diet
for 4 weeks showed no changes on glucose transport in isolated epitrochlearis muscle in
response to insulin (99).
1.3.3. The effects of ethanol on insulin signaling pathway
While the mechanisms for disrupted insulin-stimulated glucose utilization after
ethanol exposure are not well understood, ethanol disrupts the insulin signaling pathway
in a number of cell types. Moderate amounts of alcohol intake have been reported to
enhance insulin sensitivity (94;95). Coincidently, studies in rodents reveal increased
- 19 - phosphorylation of IRS-2 and PKB/Akt in the liver of rats consuming 3% ethanol for 4
weeks (100).
In contrast, excessive ethanol consumption leads to insulin resistance, regardless it is a
single or continuous intake (58;96;97). However, the effects of excessive ethanol
consumption on the insulin signaling are still inconsistent and unclear. One treatment of
a large dose of ethanol in rats increases tyrosine phosphorylation of insulin receptor, IRS-
1, and IRS-2, as well as increases PI 3-kinase activity in muscle, hepatic, and adipose tissues in response to insulin stimulation, despite the presence of insulin resistance (97).
The authors suggest that the insulin signaling step impaired by ethanol is likely to be
downstseam from PI 3-kinase.
With respect to chronic excessive ethanol exposure, phosphorylation of insulin receptor substrate-1 and activities of PI 3-Kinase and Akt/PKB are reduced in the liver and developing brain, leading to impaired cell growth and survival (101;102). In the liver,
TRB3 has been recently reported to inhibit insulin signaling by binding directly to Akt
and preventing activation of this kinase (103). Studies including liver lysates isolated
from ethanol-fed rats and hepatoma cell line FGC-4 after ethanol treatment have shown that ethanol exposure induces TRB3, which prevents Akt association with the plasma membrane, and subsequent Akt-associated signaling (104). Conversly, other studies report that insulin signaling via PI 3-kinase to PKB/Akt phosphorylation is not
suppressed by chronic ethanol in either liver, adipose tissue, or skeletal muscle (97;98).
Instead, chronic ethanol feeding disrupts insulin-mediated Cbl/TC10 signaling and actin polymerization in isolated adipocytes (105). These data suggest that excessive
- 20 - consumption of ethanol induces differential effects on the insulin signaling in a tissue-
specific and dose-dependent manner.
1.4. Experimental Models of Chronic Ethanol Consumption
Alcohol consumption has been associated with a variety of adverse pathological effects. In order to investigate the pathogenesis and prevention of these alcohol-related
pathologies, animal models of alcohol administration have been developed to mimic the
pathological changes occurring in humans. Two major experimental models in animals
have been developed, Lieber-DeCarli liquid diet technique and Tsukamoto-French
intragastric feeding of liquid diet in rodents (106). The use of these animal models
allows for the investigation of ethanol’s effects apart from the consequences of nutritional
alterations. The development, importance, and comparison of these two animal models
are discussed here.
1.4.1. Lieber-DeCarli liquid diet
The Lieber-DeCarli liquid diet technique has been used for more than 40 years in
studying many of the pathological disorders associated with alcoholism (107).
Administration of ethanol as a part of nutritionally defined liquid diet overcomes the
natural aversion of most animals to ethanol, compared to providing ethanol in the
drinking water (108). The Lieber-DeCarli diet also allows for precise control of the
nutritional intake of the animals during ethanol exposure. The standard liquid diet
contains an amount of fat (35% of total energy) comparable to that of the American diet,
18% as protein, and, in the control diet, 47% as carbohydrate. In the ethanol-containing
- 21 - diet, 36% of carbohydrate is isocalorically replaced by ethanol (Table 1.1). An adequate
content of all necessary nutrients is also included in the diet, such as various vitamins and
minerals at or above the daily recommended allowance (108). Plasma ethanol
concentrations of 20-40 mM (90-180 mg/dl) are routinely observed with this diet (109).
Feeding of animals with this liquid diet produces many of the same pathologies observed in alcoholics, including hepatic steatosis, hyperlipidemia, various metabolic and
endocrine disorders (108). When baboons are fed this diet for more than 2 years, more
severe stages of alcoholic liver disease also develop, including cirrhosis (110).
In the ethanol diet, carbohydrate accounts for only 11% of total calories. It has been
questioned whether the decreased supply of carbohydrate might be responsible for some
of the pathophysiological effects attributed to the ethanol. However, low carbohydrate
diets alone, i.e. isocalorical substitution of ethanol for fat rather than carbohydrate, do not
induce any histological change or fat accumulation in the liver (111). In addition, it is
conceivable that low levels of carbohydrate in the ethanol diet might cause a decrease in
circulating plasma insulin; however, fasting plasma insulin concentration is not different
between pair- and ethanol-fed rats (99). These data suggest that the detrimental changes
in the liver, as well as other metabolic and endocrine disorders that are observed in
ethanol-fed rats, are due to ethanol, as opposed to differences in carbohydrate content in
the diets.
The fat content of the diet (35% of total energy) is considered high for rats, but it
mimics the average American diet (109). Additionally, the high fat content in the diet is
critical for the development of fatty liver since the quantity of fat in the diet has a dose-
dependent effect on the amount of fat that accumulates in the rat liver (112). Yet, even a
- 22 - Table 1.1 Macronutrient composition of Liber-DeCarli liquid diet1.
Control Diet Ethanol Diet Fat: 35% total 35% total Corn Oil 7.3% 7.3% Olive Oil 25.2% 25.2% Safflower Oil 2.5% 2.5% Carbohydrate (maltose dextrin) 47% 11% Protein 18% 18% Ethanol 0% 36%
1% calories contribution to the diet.
- 23 - low-fat (5-10%) ethanol-containing diet can induce some of the pathologies of ethanol,
such as the induction of the microsomal ethanol-oxidizing system in rats (113) and
accumulation of hepatic triglycerides in mice (114).
1.4.2. Tsukamoto-French Rat Model
While the Lieber-DeCarli liquid diet provides a technically simple way to reproduce early stages of alcoholic liver injury, it limits the ethanol consumption in rats to 36% of total caloric intake compared to greater than 50% of ethanol intake in some alcoholics.
To overcome this limitation, Tsukamoto and French developed a model in which liquid diets are administrated by continuous intragastric infusion (115). This technique allows for substantial increase in the overall dose of ethanol administrated in rats. Plasma ethanol concentrations of 50-80 mM (230-370 mg/dl) are achieved with this approach
(116), and more advanced liver injury including inflammation and fibrosis also develops in these rats (115). Therefore, Tsukamoto-French rat model provides a valuable way to study more advanced liver injury. However, the continuous intragastric administration of excessive ethanol is not physiological because of a constant challenge of rats being in a fed state. Moreover, intragastric administration is much more cumbersome and expensive than the Lieber-DeCarli diet.
In summary, the Lieber-Decarli liquid diet technique for administrating ethanol orally
is economic and suitable for most experimental applications, particularly those
investigating the mechanisms for the development of early stages of liver damage. By
using the Tsukamoto-French rat model, one can study the mechanisms of more severe
liver damage, such as inflammation and fibrosis. In this thesis, the Lieber-DeCarli liquid
- 24 - diet has been used to investigate the effects of ethanol on lipid and glucose homeostasis in rat adipose tissue, which has been hypothesized to contribute to the pathogenesis of early stage of liver damage caused by ethanol.
- 25 - Chapter 2
Research Hypotheses and Objectives
2.1. Introduction
Chronic ethanol consumption is associated with insulin resistance and is considered as
an independent risk factor for the development of type 2 diabetes. Although the exact
mechanisms by which chronic ethanol induces these pathophysiological conditions are
not well understood, the disruption of lipid and glucose homeostasis may contribute to
the development of chronic ethanol-induced pathology. Adipose tissue, as the major
storage site of fat, as well as one of the major sites for glucose transport, plays an
important role in maintaining whole-body lipid and glucose homeostasis. Importantly,
insulin regulates both lipid and glucose metabolism in adipose tissue, which puts the high
importance on the sensitivity of adipose tissue to insulin with respect to lipid and glucose
homeostasis since insulin resistance is associated with many diseases including obesity and type 2 diabetes. Due to the prevalence of type 2 diabetes and the poor understanding of the interaction of chronic ethanol consumption with type 2 diabetes, it is important to
study the effects of chronic ethanol exposure on lipid and glucose homeostasis in adipose tissue, as well as the effects of chronic ethanol on the hormonal regulation of these pathways, including regulation by β-adrenergic receptor agonists and insulin.
2.2. Research Hypotheses and Objectives
Chronic ethanol feeding disrupts both G protein- and insulin-dependent signal
transduction, which are the signaling pathways involved in the regulation of both lipid
- 26 - and glucose metabolism in adipose tissue. We hypothesized that the regulation of lipid
and glucose metabolism in adipose tissue might be vulnerable to long-term exposure to
ethanol. Chronic ethanol feeding to rats increases circulating free fatty acid concentrations (60). Since lipolysis in adipose tissue mobilizes free fatty acids to the circulation, we therefore hypothesized that chronic ethanol feeding increased lipolysis in adipose tissue. Moreover, since both G protein- and insulin-dependent signal pathways, which have been shown affected by chronic ethanol exposure, regulate lipolysis, we hypothesized that increased lipolysis resulted from a disruption of hormonal regulation by ethanol.
Chronic ethanol feeding decreases whole-body glucose utilization during the
hyperinsulinemic-euglycemic clamp (58;97). Adipose tissue and skeletal muscle are the
two major sites for glucose utilization. However, the relative contribution of adipose tissue and skeletal muscle in mediating the impaired insulin-stimulated glucose disposal after chronic ethanol are unknown. As previous studies have shown that chronic ethanol exposure decreases insulin-stimulated glucose uptake in isolated adipocytes (59;98), but not in isolated epitrochlearis muscle (99), we hypothesized that chronic ethanol feeding decreased glucose disposal in adipose tissue in vivo, but may not in skeletal muscle.
The hypotheses and objectives of this study were as follows:
Hypothesis 1: Chronic ethanol feeding disrupts whole-body lipid homeostasis by
disrupting lipid metabolism in adipose tissue.
Objective 1: To investigate the effects of chronic ethanol on triglyceride turnover
2 rates in adipose tissue by the use of H2O.
- 27 - Objective 2: To investigate the effects of chronic ethanol on hormonal regulation
of adipocyte lipolysis.
Hypothesis 2: Chronic ethanol feeding disrupts insulin-stimulated whole-body glucose
utilization by impairing glucose disposal in adipose tissue.
Objective 3: To investigate the effects of chronic ethanol on tissue-specific
glucose disposal.
- 28 - Chapter 3
Dysregulation of Triglyceride Metabolism in Adipose Tissue by Chronic Ethanol
3.1. Introduction
Alcohol consumption is causally related to more than 60 different medical conditions, including hepatic diseases, cardiovascular disorders, as well as diabetes mellitus (9). In humans, chronic ethanol exposure causes excessive lipid accumulation in liver with the eventual development of hepatic steatosis (60). These pathophysiological effects of ethanol can be modeled in rodents fed diets containing ethanol; chronic ethanol feeding to rats induces hepatic steatosis coupled with the development of hyperlipidemia, characterized by elevated plasma cholesterol and triglyceride concentrations (60). These data suggest that the disruption of lipid homeostasis by ethanol is likely a mediator of alcohol-related disease progression. However, the effects of chronic ethanol feeding on lipid metabolism in adipose tissue, the biggest storage pool of lipids, are unknown.
Adipose tissue is a specialized connective tissue that functions as the major storage
site for fat in the form of triglycerides. Serving as an energy reserve, adipose tissue
synthesizes triglycerides when energy intake exceeds energy output. During fasting or in
response to infection and inflammation, adipose tissue mobilizes free fatty acids and
glycerol providing other tissues with metabolites and energy substrates (33).
Mobilization of fatty acids and glycerol from triglycerides in adipose tissue, also termed
lipolysis, is tightly regulated by a number of hormones. The primary hormones
regulating lipolysis are catecholamines, which initiate lipolysis by the stimulation of β-
adrenergic receptors, and insulin, which inhibits catecholamine-induced lipolysis (117).
- 29 - Chronic ethanol exposure disrupts both G protein- and insulin-dependent signal transduction in a variety of cell types, including adipocytes (48;101;102;105). For example, we have demonstrated that ethanol feeding for 4 wks decreases β-adrenergic receptor-stimulated lipolysis (118), and suppresses insulin-stimulated glucose uptake in isolated adipocytes (59;98). However, the effect of chronic ethanol feeding on the anti- lipolytic response of adipocytes to insulin has not been examined.
In this study, we investigated the effects of chronic ethanol feeding over a 2 wk period on the integrated rates of in vivo triglyceride turnover in rat epididymal adipose tissue by
2 2 2 the use of H2O (119). Following the administration of H2O, H in body water equilibrates with the carbon-bound hydrogens of glycerol 3-phosphate and the rates of triglyceride synthesis and degradation are determined by measuring the incorporation/ washout of 2H to/from carbon 1 of triglyceride-bound glycerol (120). The rates of triglyceride synthesis and degradation during 2 wks of ethanol feeding were both increased. Because increased rates of lipolysis are associated with the development of insulin resistance and fatty liver in other model systems, we next investigated the mechanisms by which chronic ethanol increased triglyceride degradation. We find that the ethanol-induced increased in lipolysis in adipose was due to a loss of the anti-lipolytic actions of insulin, rather than an increase in stimulation of lipolysis by β-adrenergic receptor activation.
3.2. Materials and Methods
3.2.1. Materials
- 30 - Male Wistar rats (150-160g) were purchased from Harlan Sprague Dawley
(Indianapolis, IN). The Lieber-DeCarli high-fat ethanol diet was purchased from Dyets
(Bethlehem, PA). Maltose dextrins were obtained from BioServ (Frenchtown, NJ).
Ethanol-L3K assay kit was purchased from Diagnostic Chemicals Limited (Oxford, CT).
2 2 H2O (99.9 atom percent excess) and [ H5]glycerol (98 atom percent excess) were
purchased from Isotec (Miamisburg, OH), ion-exchange resins were from Bio-Rad
(Hercules, CA), glycerokinase was from Roche (Indianapolis, IN), bis(trimethylsilyl)trifluoroacetamide with 10% trimethylchlorosilane was from Regis
Technologies Inc. (Morton Grove, IL), gas chromatography-mass spectrometry (GC-MS) supplies were from Agilent Technologies (Wilmington, DE). NEFA C Kit for the measurement of plasma free fatty acid concentration was purchased from Wako
Chemicals USA, INC. (Richmond, VA). Blood glucose meter and blood glucose test
strips were from CVS (Woonsocket, RI), human insulin was from Eli Lilly (Indianapolis,
IN), rat insulin ELISA was from Mercodia Inc. (Winston Salem, NC), and all other reagents were from Sigma (St. Louis, MO).
3.2.2. Animal protocol for determination of triglyceride turnover rates
Rats were allowed free access to the Lieber-DeCarli liquid diet containing ethanol as
35% of total calories or pair-fed an iso-caloric control diet which substituted maltose
dextrins for ethanol for 4 wks as previously described (118). Rats were housed in
individual wire-bottom cages under controlled temperature and humidity with 12 h light-
12 h dark (7:00 PM-7:00 AM) cycle. After 2 wks of feeding, rats were given an
2 2 intraperitoneal injection of H2O-saline (0.9 g NaCl in 1,000 ml 99.9% H2O, 16.25 μl/g
- 31 - 2 2 body weight). H2O was then included in the diets (enriched to 5% of H) for 5 days;
2 after that, H2O was switched to tap water. Rats were euthanized on 0, 3, 5, 7, 9, 11 and
2 14 days after the intraperitoneal injection of H2O (n = 2 per day per group except that
day 0 where n = 3). Rats were anesthetized by intraperitoneal injection of 0.075 ml for
ethanol-fed rats or 0.12 ml for pair-fed rats per 100 g body weight of a cocktail
containing 10 mg/ml acepromazine, 100 mg/ml ketamine and 20 mg/ml xylazine. The
lower dose of anesthetic for ethanol-fed rats was used because of an increased sensitivity
to the anesthetic cocktail after ethanol feeding. However, it is unlikely that the different
doses had any impact in our assay, as ketamine and xylazine at these doses either have no
effects or equivalent effects on lipogenic and lipolytic activities of adipose tissue
(121;122). Under anesthesia, blood was collected and epididymal adipose tissue was
frozen in liquid nitrogen and stored at -80˚C. Plasma samples were prepared by centrifugation at 16,100 x g for 2 min and plasma ethanol concentration was measured
immediately by the Ethanol-L3K kit. The rats used in these studies were not fasted; all
studies were carried out at 10:30 AM, except the hyperinsulinemic-euglycemic clamps
which were performed at 12:00 noon (as time 0 of the clamps). Procedures involving
animals were approved by the Institutional Animal Care and Use Committee at Case
Western Reserve University.
3.2.3. Measurements of isotope enrichment by gas chromatography-mass spectrometry
The 2H-labeling of body water-The 2H-labeling of body water was assayed by
exchange with acetone as described by Yang et al. (123) and as modified previously
(119;120). Briefly, known 2H atom percent excess standards were prepared by mixing
- 32 - 2 naturally labeled water and 99.9% H2O. Assays were performed using 40 μl of plasma
or standard, 2 μl of 10 N NaOH, and 4 μl of a 5% (v/v) solution of acetone in acetonitrile.
After overnight incubation, the solution was extracted with 600 μl of chloroform and
2 dried with Na2SO4. The H-labeling of acetone was then determined by GC-MS. Ions of
mass-to-charge ratios (m/z) 58-60 were monitored.
The 2H-labeling of triglyceride-bound glycerol-Frozen epididymal adipose tissue was
hydrolyzed with 1 N KOH in 90% ethanol at 70˚C for 2 hrs. After evaporation of ethanol, free glycerol was recovered as previously described (119;120). H2O (3 ml) was added
and the solution was acidified to ~pH 1. After extraction of fatty acids by diethyl ether (3
x with 4 ml), the aqueous solution was neutralized to ~pH 7, and free glycerol in the
aqueous solution was reacted with 0.2 M ATP and 5 U of glycerokinase at 37˚C for 2.5 hrs. The formed glycerol 3-phosphate was then purified by passing the solution over an
AG 1-X8 resin (formate form), washing the column with water, and eluting the column with 4 N formic acid. The labeling of 2H bound to carbon 1 of glycerol 3-phosphate was
determined by reacting the glycerol 3-phosphate with 100 µl of
bis(trimethylsilyl)trifluoroacetamide + 10% trimethylchlorosilane for 30 min at 75°C.
Isotope enrichment was determined by GC-MS. Ions of mass-to-charge ratios (m/z) 445-
446 and 357-358 were monitored.
3.2.4. Mathematical model for determination of triglyceride turnover rates
The rates of triglyceride synthesis and degradation were determined by fitting the
concentration of triglyceride-bound glycerol (Fig 3.2B) and 2H-labeling of body water and triglyceride-bound glycerol (Fig 3.4) as previously reported (119;120). Briefly, the
- 33 - concentration of triglyceride-bound glycerol was first modeled. The incorporation of 2H
from body water into lipids was then modeled using a single-compartment model,
assuming that the labeling of plasma water reflects that of water in adipose tissue. The
parameters of interest, the rates of triglyceride synthesis and degradation, were then
estimated from the data by using nonlinear least-squares fitting.
3.2.5. Ethanol elimination from blood in rats
After 4 weeks of ethanol feeding, rats were transported from the Animal Resource
Center at 7:00 AM in the morning. Blood (100 µl) was collected from the rat tail vein every hour. Plasma was separated by centrifugation for 2 min at 16,100 x g and plasma ethanol concentration was measured immediately using the Ethanol-L3K assay kit.
2 3.2.6. Determination of body fat mass by H2O dilution space
After 4 weeks of pair- or ethanol-feeding, rats were weighed and received an
2 2 intraperitoneal injection of 2.5 ml of H2O-saline (0.9g NaCl in 1,000ml 99.9% H2O) to establish 1-1.5% 2H enrichment of body water. After 3 hours, rats were transported from
the Animal Resource Center and anaesthetized. Blood (~1ml) was collected from the
portal vein in EDTA tubes and plasma was separated by centrifugation for 2 min at
16,100 x g. The 2H labeling of plasma was then measured as before. Body fat mass was
2 calculated by the H2O dilution space as previously described (124).
3.2.7. β-adrenergic receptor-stimulated in vivo lipolysis
- 34 - After 4 wks of pair- or ethanol-feeding, rats were anesthetized by inhalation of an
isoflurane and oxygen mixture. Under anesthesia, rats were given an intraperitoneal
injection of the β3-adrenergic receptor agonist, CL316,243 at a dose of 0.1 mg/kg body
weight. Blood samples were collected before and at 8, 16, 30, and 60 min after the
injection from the tail vein. Plasma was separated and plasma glycerol and free fatty acid
concentrations were then determined as indices of lipolysis.
3.2.8. Isolation of adipocytes and ex vivo lipolysis assay
After 4 wks of feeding, rats were anesthetized by an intraperitoneal injection of a
cocktail as described above and epididymal adipose tissue was removed. Ex vivo
lipolysis was measured as glycerol released into the cell medium over 1 hour as described before (118). Briefly, adipocytes were isolated by collagenase digestion (118), and cell concentration was adjusted to 1 x 106 cells/ml. 200 μl aliquots of cells were placed into 5
ml polypropylene tubes and 1 µM isoproterenol, a β-adrenergic receptor agonist, and/or
increasing concentrations of insulin were added. Adipocytes were incubated for 1 hour at
37˚C in a shaking water bath (100 rpm), and glycerol concentration in the cell medium
was measured using Free Glycerol Reagent (GPO trinder reagent).
3.2.9. Hyperinsulinemic-euglycemic clamp and the appearance rate of glycerol (Ra)
Hyperinsulinemic-euglycemic clamp-After 3 wks of pair- or ethanol-feeding, rats were
anesthetized by inhalation of an isoflurane and oxygen mixture and the left carotid artery
and the right jugular vein were catheterized for blood sampling and intravenous infusion
during the clamp, respectively. All rat surgeries were done in the Mouse Metabolic and
- 35 - Phenotyping Center at Case Western Reserve University. Rats were allowed a week to
recover from the surgery while maintained on their respective diets. Hyperinsulinemic-
euglycemic clamps were then performed on one rat at a time as previously described
(125), with minor modifications. Briefly, rats were transported from the Animal
Resource Center and allowed at least 90 min to stabilize before commencement of the
2 glucose clamp. [ H5]Glycerol (~1 µmol/kg/min) was continuously infused from the
jugular vein catheter for 90 min basal period and 2-h clamp period. Baseline levels of
blood glucose, plasma insulin, plasma glycerol and free fatty acid, and the plasma
appearance rate of glycerol (Ra) were determined as the mean of values obtained in blood
samples collected at -30 and -5 min. At time 0, a primed (60 mU/kg)/continuous (4
mU/kg/min) infusion of human insulin was started and continued for 2 hrs. The blood
glucose concentration was clamped at euglycemic level by a variable rate infusion of
20% glucose. Blood glucose was monitored with a blood glucose meter and the rate of glucose infusion adjusted every 10 min. Blood samples for determination of plasma
2 insulin, plasma glycerol and free fatty acid concentrations, and plasma [ H5]glycerol
enrichments were obtained at -30, -5, 90, 100, 110, and 120 min. There were no
differences in the hematocrit between pair- and ethanol-fed rats either before or after the
clamp (Pair-fed: from 35 ± 2% to 30 ± 2%; Ethanol-fed: from 35 ± 5% to 26 ± 3%).
Appearance rate of glycerol in plasma (Ra)- The plasma glycerol Ra was used as an index for systemic lipolysis, as calculated by the following equation: Ra =
(ENRinf/ENRpl-1) · F, where ENRinf is the isotopic enrichment of the infusate, ENRpl is
the isotopic enrichment of plasma and F is the rate of the isotope infusion (126). The 2H-
labeling of plasma glycerol was determined as described below. 20 µl of plasma was
- 36 - deproteinized with 200 µl of methanol by centrifugation for 10 min at 16,100 x g. The
fluid fraction was then evaporated to dryness and reacted with 50 µl of bis(trimethylsilyl)trifluoroacetamide + 10% trimethylchlorosilane for 20 min at 75°C.
Isotope enrichment was determined by GC-MS. Ions of mass-to-charge ratios (m/z) 205-
208 were monitored.
3.2.10. Statistical analyses
Data are expressed as means ± SD where n = 2, and as means ± SEM where n ≥ 3.
Dose-response curves to insulin in isolated adipocytes were estimated by non-linear
regression (GraphPad Prism® 4; San Diego, CA). Statistical analyses were performed
using the general linear model procedure on SAS for personal computers. Differences
between groups were determined by least square means.
3.3. RESULTS
3.3.1. Characteristics of ethanol-fed rats
To study the effects of chronic ethanol feeding on lipid metabolism in adipose tissue,
rats were fed a liquid diet with or without ethanol for 4 wks. Rats gained weight over
time on both pair- and ethanol-liquid diets, and there were no differences in body weights
between pair- and ethanol-fed rats either before or after feeding for 2-4 wks (Table 3.1).
Plasma ethanol concentration measured at 10:30AM, was 15-17mM in ethanol-fed rats, and non-detectable in pair-fed rats (Table 3.1).
- 37 - Table 3.1 Rat body weights, plasma ethanol concentrations and epididymal fat weights
Weeks on diet 0 2 3 4 Pair-fed 168 ± 8 234 ± 5 259 ± 7 272 ± 5 Body weight (g) EtOH-fed 186 ± 6 237 ± 6 255 ± 6 280 ± 6 Pair-fed --- N.D. N.D. N.D. Plasma ethanol (mM) EtOH-fed --- 15.3 ± 1.9 16.6 ± 1.2 14.9 ± 1.6 Epididymal fat Pair-fed --- 2.7 ± 0.1 4.1 ± 0.2 4.8 ± 0.4 weight (g) EtOH-fed --- 2.1 ± 0.2* 2.9 ± 0.2* 3.3 ± 0.1*
Data are expressed as mean ± SD for plasma ethanol data (n = 2), and as mean ± SEM for body weight data (n = 13) and epididymal fat weight data (n = 7 for wk 2, n = 6 for wk 3, and n = 4 for wk 4). N.D. = not detectable. *p < 0.05 Pair-fed vs. EtOH-fed.
- 38 - To further determine how fast ethanol eliminates from the rats after the removal of
diet, we determined plasma ethanol concentrations at different times during the day. In
the early morning at 7:00 AM, plasma ethanol concentration was as high as 50 mM, and
it declined rapidly. At 12:00 noon, plasma ethanol concentration was close to zero (Fig
3.1). The half-life of plasma ethanol was about 1.5 hours.
Epididymal fat weight increased over time in both pair- and ethanol-fed rats; the rate
of increase was not affected by chronic ethanol feeding (Fig 3.2A). However, epididymal
adipose tissue weight was lower in rats after ethanol feeding for 2, 3, and 4 weeks
compared to pair feeding (Table 3.1). These data suggest that the rates of epididymal fat
accumulation over time in rats did not differ between pair- and ethanol-feeding for 2-4
weeks, although the weights were already lower in ethanol-fed rats (Table 3.1).
Consistent with the decrease in weights of epididymal adipose tissue, the total amounts of triglyceride-bound glycerol isolated from epididymal fat were also lower in ethanol-fed rats compared to pair-fed rats based on the linear regression analyses (Fig 3.2B).
Since the lipid content in epididymal adipose tissue was decreased in rats after ethanol
feeding, we next asked whether the total body fat mass was also decreased by chronic
2 ethanol feeding. The percentage of body fat was calculated from the H2O dilution space
(124). There was no difference in the percentage of total body fat between pair- and
ethanol-fed rats (Fig 3.3).
3.3.2. Triglyceride turnover rates
To further investigate the effects of ethanol on metabolic activity of adipose tissue, the
rates of triglyceride synthesis and degradation in epididymal adipose tissue were
- 39 - 50 M)
m 40
30
aethanol( 20 m
Plas 10
0 7 8 9 10 11 12 13 14 15 Time of the day (hr)
Figure 3.1 Plasma ethanol decay in rats chronically fed with ethanol for 4 weeks.
Blood was collected from the tail vein of rats after ethanol feeding for 4 weeks. Plasma ethanol concentration was measured by the Ethanol-L3K kit. Data are expressed as mean
± SD (n = 2).
- 40 - AB
D OHO 7 D2OH2O 1400 2 2
6 1200
5 1000
4 800
3 600 moles/fat) μ ( 2 400 2 2 Pair-fed r =0.73 Pair-fed r =0.76 2
Fat pads (g1 wet weight) 200 2 EtOH-fed r =0.68 EtOH-fed r =0.71 Triglyceride-bound glycerol 0 0 2 3 4 2 3 4 Weeks on diet Weeks on diet
Figure 3.2 Chronic ethanol feeding decreased lipid content in epididymal adipose
tissue.
A) The changes in epididymal adipose tissue wet weight over time were analyzed by
linear regression. Pair-fed: y = 0.886x + 1.126, r2 = 0.76; EtOH-fed: y = 0.497x + 1.169,
r2 = 0.71. B) A known amount of epididymal adipose tissue was hydrolyzed with 3 mL
of 1 N KOH in 90% ethanol at 70˚C for 2 hrs. Glycerol concentration was then determined. Data were analyzed by linear regression. Pair-fed: y = 237.1x + 18.1, r2 =
0.73; EtOH-fed: y = 185.7x + 63.5, r2 = 0.68. Each data point represents the mean ± SD
(n = 2 per day per group except wk 2 where n = 3). The general linear model procedure
on SAS was used to statistically analyze the linear regression of the data. p < 0.05 for the y intercepts of two lines within each panel at x = 2 wks; the slopes of the lines did not differ between pair- and ethanol-fed rats.
- 41 -
25
20
15
10
Body fat mass (%) fat mass Body 5
0 Pair-fed EtOH-fed
Figure 3.3 Body fat mass in pair- and ethanol-fed rats.
2 After 4 weeks of feeding, rats were given an intraperitoneal injection of H2O-saline.
Blood was collected from the portal vein 3 hours after the injection. The 2H-labeling of plasma was then measured and the percentage of total body fat was calculated from the
2 dilution space of H2O. Data are expressed as mean ± SEM (n = 6).
- 42 - 2 determined by the use of H2O as described in MATERIALS AND METHODS
(119;120). Figures 3.4A and 3.4B show the 2H-labeling curves of body water in pair- and
2 ethanol-fed rats. The initial intraperitoneal injection of H2O enriched body water to approximately 2.1 molar percent excess in both pair- and ethanol-fed rats. Five-day
2 maintenance on H2O in the diets increased the labeling to 3.4 molar percent excess.
2 2 When rats were switched from H2O to tap water, H was washed out of body water. The
calculated t1/2 of body water did not differ between pair- (2.1 days) and ethanol-fed (2.2
days) rats (Fig 3.4 A/B).
To determine the kinetics of triglyceride turnover, the labeling of 2H bound to C1 of
triglyceride-glycerol was measured in epididymal adipose tissue from both pair- and
2 ethanol-fed rats. While rats were maintained on H2O, the labeling of triglyceride-
glycerol increased in both groups; after switching to tap water, the labeling of triglyceride-glycerol decreased (Fig 3.4 C/D). The calculated rates of triglyceride
turnover show that 4-week ethanol feeding increased both the rates of triglyceride
synthesis and degradation (Table 3.2). However, the net accumulation rates of
triglycerides in epididymal adipose tissue were not different between two groups (Table
3.2), which is consistent with the unchanged slopes of linear regression of epididymal fat
content (Fig 3.2). Taken together, these data suggest that ethanol feeding for 2-4 wks
stimulated the metabolic activity of epididymal adipose tissue.
3.3.3. β3-adrenergic agonist-stimulated in vivo lipolysis
Hydrolysis of triglycerides in adipocytes is primarily regulated by the activity of the
sympathetic nervous system and by plasma insulin level (117). Catecholamines stimulate
- 43 - Pair-fed EtOH-fed
A D OHO B D OHO 4.0 2 2 4.0 2 2 3.5 3.5 3.0 3.0 2.5 2.5 2.0 2.0 1.5 1.5 1.0 1.0 (moler percent excess ) (molar percent excess ) H-labeling of body water H-labeling of body water
2 0.5 2 0.5 0.0 0.0 2 3 4 2 3 4 Weeks on diet Weeks on diet C D 2.5 2.5
2.0 2.0
1.5 1.5
1.0 1.0
0.5 0.5 H-labeling of triglyceride-bound H-labeling of triglyceride-bound glycerol (molar percent excess ) glycerol (molar percent excess ) 2 0.0 2 0.0 2 3 4 2 3 4 Weeks on diet Weeks on diet
Figure 3.4 2H-labeling of body water and triglyceride-bound glycerol isolated from
epididymal adipose tissue in rats.
A, B) Labeling of body water was assayed by the method of hydrogen exchange with acetone in plasma. C, D) Frozen epididymal adipose tissue was hydrolyzed with 3 mL of
1 N KOH in 90% ethanol at 70˚C for 2 hrs to isolate triglyceride-bound glycerol. The labeling of 2H bound to carbon 1 of triglyceride-glycerol was determined as described in
MATERIALS AND METHODS. Mathematical model described in MATERIALS AND
METHODS were used to fit the 2H-labeling of triglyceride-bound glycerol. Each data
point represents the mean ± SD (n = 2 per day per group except wk 2 where n = 3).
- 44 - Table 3.2 Rates of triglyceride synthesis and breakdown.
Triglyceride kinetics Triglyceride synthesis Triglyceride Net Diet (µmol/day) degradation (µmol/day) (µmol/day) Pair-fed 43.7 ± 3.6 15.2 ± 3.6 28.5 EtOH-fed 81.5 ± 11.7 47.0 ± 11.7 34.5
The rates of triglyceride synthesis and degradation were determined by fitting the
concentration of triglyceride-bound glycerol (Fig 3.2B) and 2H-labeling of body water and triglyceride-bound glycerol (Fig 3.4) using the mathematical model described in
MATERIALS AND METHODS.
- 45 - lipolysis by activating β-adrenergic receptors, while insulin acts as a counter-regulator to inhibit lipolysis through the insulin receptor (117). Since chronic ethanol feeding increased the rate of triglyceride degradation in epididymal adipose tissue, we hypothesized that ethanol activated the lipolytic action of β-adrenergic receptors and/or suppressed the anti-lipolytic action of insulin. We have recently reported that chronic ethanol feeding decreases β-adrenergic receptor-stimulated lipolysis in isolated adipocytes (118). To validate these ex vivo experiments in the intact animals, we first measured in vivo lipolysis stimulated by CL 316,243, an agonist specific for β3- adrenergic receptors, the primary isoform of β-adrenergic receptors expressed in adipose tissue (127). Baseline concentration of plasma glycerol and free fatty acids were not different between pair- and ethanol-fed rats (Fig 3.5). Intraperitoneal injection of CL
316,243 increased plasma glycerol and free fatty acid concentration in both pair- and ethanol-fed rats. However, the β3 agonist-mediated elevation of plasma glycerol and free fatty acid concentrations was suppressed in ethanol-fed rats compared to pair-fed rats
(Fig 3.5). These data indicated that chronic ethanol feeding to rats decreased β- adrenergic receptor-stimulated lipolysis, suggesting that the increase in the rate of triglyceride degradation observed in vivo in chronically ethanol-fed rats was not mediated by an increased lipolytic response of adipocytes to β-adrenergic activation.
3.3.4. Anti-lipolytic action of insulin ex vivo
In contrast to the role of β-adrenergic agonist in the activation of lipolysis, insulin is the key inhibitor of lipolysis. Since insulin signaling pathways are also targets of chronic ethanol, we next examined the effect of chronic ethanol on insulin-mediated inhibition of
- 46 - A Pair-fed EtOH-fed 750
600 M) μ
450
300 * * * 150 * Plasma glycerol (
0 0 10 20 30 40 50 60
Time afte r β 3 agonist administration (min)
B Pair-fed 1.75 EtOH-fed 1.50
1.25
1.00 * * 0.75 * 0.50 * Plasma FFA (mM) 0.25
0.00 0 10 20 30 40 50 60 Time afte r β 3 agonist administration (min)
Figure 3.5 Four-week ethanol feeding decreased β3-adrenergic receptor agonist,
CL316,243-stimulated systemic lipolysis.
After 4 wks of pair- or ethanol-feeding, rats were anesthetized and received an intraperitoneal injection of an agonist of β3-adrenergic receptors, CL316,243 at a dose of
0.1mg/kg body weight. Blood samples were collected before and at 8, 16, 30, and 60 min after the injection from the tail vein. Plasma glycerol (A) and free fatty acid (FFA) (B) concentrations were then determined as indices of lipolysis. Data are expressed as mean
± SEM (n = 6). *p < 0.05 Pair-fed vs. Ethanol-fed.
- 47 - lipolysis in an ex vivo model of isolated adipocytes. Consistent with our previous study
(118), ethanol feeding for 4 wks decreased isoproterenol-stimulated lipolysis in isolated adipocytes, without changing basal rates of lipolysis (Fig 3.6A). Insulin, at concentrations from 0.1-100 nM, dose-dependently decreased isoproterenol-stimulated lipolysis in adipocytes isolated from pair-fed rats (Fig 3.6). In contrast, at these physiological concentrations, insulin did not inhibit lipolysis in adipocytes isolated from ethanol-fed rats (Fig 3.6). Maximal inhibition of isoproterenol-stimulated lipolysis was observed with 10 µM insulin and did not differ between adipocytes from both pair- and ethanol-fed rats (Fig 3.6A).
3.3.5. Anti-lipolytic action of insulin in vivo
To validate the ex vivo effects of chronic ethanol on the sensitivity of lipolysis to insulin in an in vivo model, we utilized the hyperinsulinemic-euglycemic clamp technique
(126). Plasma insulin levels were maintained at a steady state in both pair- and ethanol- fed rats during 90-120 min of the clamp. Glucose infusion rate during this time period was also constant (data not shown). Therefore, the last 30 min of the clamp (90-120 min) was assumed to be at steady state and the means of the 90, 100, 110, and 120 min values were used for steady-state values (126).
Body weight of rats used in the hyperinsulinemic-euglycemic clamps did not differ
between pair- and ethanol-fed rats (Table 3.3). There were no differences between pair- and ethanol-fed rats in basal blood glucose, mean blood glucose at 90-120 min of the glucose clamp, mean plasma insulin levels at 90-120 min achieved during insulin
- 48 - A B Pair-fed Pair-fed 1.00 1.4 EtOH-fed * EtOH-fed * 1.2 0.75 1.0
+ * 0.8 0.50 + + * * 0.6 0.4 Glycerol release 0.25 Glycerol release ** 0.2 (fold of own Iso-stimulated)
(fold Iso-stimulated of pair-fed) 0.00 0.0 Iso (μM) 0 1 1 1 1 1 1 1 0.00.1110100 Ins (nM) 0 0 0.1 1 10 50 100 10 μM Insulin (nM)
Figure 3.6 Chronic ethanol feeding impairs the ability of insulin to inhibit lipolysis in isolated adipocytes.
A) Adipocytes isolated from epididymal adipose tissue in rats fed with pair- or ethanol- diets for 4 wks were treated with or without 1 μM isoproterenol (Iso) and increasing concentrations of insulin (Ins). Ex vivo lipolysis was determined as glycerol release over
1 hour. Data were normalized to isoproterenol-stimulated glycerol release in pair-fed group (1.05 ± 0.09 μmol/106 cells). B) Data were normalized to isoproterenol-stimulated glycerol release of each group (Pair-fed, 1.05 ± 0.09 μmol/106 cells; Ethanol-fed, 0.41 ±
0.07 μmol/106 cells) and shown as means ± SEM (n = 4 or 5). Non-linear regression
(curve fit) was used to generate inhibition curves. *p < 0.05 compared to adipocytes
treated with isoproterenol within each diet group; +p < 0.05 Pair-fed vs. EtOH-fed.
- 49 - Table 3.3 Characteristics of rats in hyperinsulinemic-euglycemic clamp study
Pair-fed EtOH-fed Numbers of rats 8 7 Body weight (g) 275 ± 5 256 ± 10 Plasma glycerol (μM) Baseline 97 ± 10 95 ± 16 Plasma FFA (mM) Baseline 0.51 ± 0.03 0.49 ± 0.06 Baseline 5.8 ± 0.3 6.1 ± 0.7 Blood glucose (mM) 90-120 min 5.3 ± 0.5 6.2 ± 1.0 Baseline 234 ± 34 117 ± 22* Plasma insulin (pmol/l) 90-120 min 534 ± 41 488 ± 32
Data are expressed as mean ± SEM. *p < 0.05 Pair-fed vs. EtOH-fed.
- 50 - infusion (Table 3.3). However, basal plasma insulin was lower in rats after 4-week
ethanol feeding compared to pair feeding (Table 3.3).
To test the sensitivity of systemic lipolysis to insulin, we first measured plasma
glycerol and free fatty acid concentrations during 90-120 min of the clamps. Basal plasma glycerol and free fatty acid concentrations were not different between pair- and ethanol-fed rats (Table 3.3). In response to insulin, plasma glycerol concentration was decreased by 56% from basal level in pair-fed rats compared to only 20% in ethanol-fed rats at the steady state (Fig 3.7 A/B). Plasma free fatty acid concentrations were decreased by 64% in pair-fed rats compared to 36% in ethanol-fed rats at the steady state
(Fig 3.7 C/D). Plasma glycerol Ra was then determined as an index for systemic lipolysis (126). Basal glycerol Ra did not differ between pair- and ethanol-fed rats; however, at the steady state of the clamp, plasma glycerol Ra was decreased by 28% in pair-fed rats with no change in ethanol-fed rats (Fig 3.8). These data suggest that chronic ethanol feeding impaired the ability of insulin to inhibit systemic lipolysis, which may contribute to the increased rate of triglyceride degradation observed in vivo after ethanol exposure.
3.4. Discussion
Chronic ethanol consumption disrupts lipid homeostasis in the liver, as well as at the
whole-body level, in both humans and animal models (60). While it is clear that the regulation of lipid homeostasis in adipose tissue plays an important role in maintaining whole-body lipid homeostasis, the effects of chronic ethanol on lipid metabolism in adipose tissue have not been studied. Here we report that chronic ethanol feeding to rats
- 51 - A B Pair-fed 1.2 EtOH-fed 1.00 * 1.0
0.8 0.75
0.6 * * + * 0.50 0.4
0.25 (fold of own basal) a glycerol (fold0.2 of own basal) m 0.0 Plas 0 90 100 110 120 0.00
Mean plasma glycerol at 90-120min Pair-fed EtOH-fed Time of the insulin clamp (min)
C D
1.2 Pair-fed 0.8 EtOH-fed * 1.0 0.7
0.8 0.6 0.5 0.6 * 0.4 0.4 + 0.3
a FFA (fold of own basal) 0.2 (fold of own basal)
m 0.2 0.1
Plas 0.0 0 90 100 110 120 Mean plasma FFA at 90-120min 0.0 Pair-fed EtOH-fed Time of the insulin clamp (min)
Figure 3.7 Plasma glycerol and free fatty acid concentrations during the
hyperinsulinemic-euglycemic clamp.
Hyperinsulinemic-euglycemic clamps were performed as described in MATERIALS
AND METHODS. A, C) Plasma glycerol and free fatty acid (FFA) concentrations during
the hyperinsulinemic-euglycemic clamp were normalized to basal concentrations of
plasma glycerol and free fatty acids within each group. B, D) Mean plasma glycerol and
free fatty acid (FFA) concentrations were shown as the means of 90, 100, 110, and 120 min values as the steady state values. Data represents the mean ± SEM (n = 7 for pair- fed group and 6 for ethanol-fed group). *p < 0.05, +p = 0.07 Pair-fed vs. EtOH-fed.
- 52 - A Pair-fed B Pair-fed EtOH-fed
in) 14 15 EtOH-fed a a m a +++ 12 + ol/kg/ 10 b m 10 * 8 * * aglycerolRa * m 6 mol/kg/min)
5 μ ( 4
aglycerolRa(u 2 m Mean plas 0 0 Plas 0 90 100 110 120 Baseline Insulin clamp Time of the insulin clamp (min)
Figure 3.8 Chronic ethanol feeding inhibits the anti-lipolytic response of adipocytes to insulin in vivo.
Plasma glycerol Ra, at the steady state during 90-120 min of the hyperinsulinemic- euglycemic clamp, was measured as an index for systemic lipolysis. A) Plasma glycerol
Ra was calculated as described in MATERIALS AND METHODS. B) Mean plasma glycerol Ra was shown as the mean of 90, 100, 110, and 120 min values as the steady state values. Data represents the mean ± SEM (n = 7 for pair-fed group and 6 for ethanol-fed group). *p < 0.05 compared with own basal within each group; +p < 0.05
Pair-fed vs. EtOH-fed. Values with different letters are significantly different (p < 0.05).
- 53 - increased the turnover rates of triglycerides, increasing both the rate of triglyceride
synthesis and degradation in epididymal adipose tissue. While the mechanisms by which
chronic ethanol increased the rate of triglyceride synthesis are still unknown, here we
have found that the increase in triglyceride degradation during ethanol feeding was
associated with a loss in insulin-mediated inhibition of lipolysis. Triglyceride
degradation was assessed both ex vivo and in vivo, including the utilization of isolated
primary adipocytes, hyperinsulinemic-euglycemic clamp technique, as well as the 2-week
2 integrated measurement of triglyceride turnover in rats by the use of H2O. Each of
these different measurements of triglyceride degradation consistently demonstrated that
chronic ethanol feeding disrupted the regulation of adipose tissue metabolism; the net
impact of these changes led to an increased lipolytic activity in adipose during ethanol
exposure.
2 H2O is used to quantify a number of biochemical parameters in vivo, including rates
of carbohydrate, protein, lipid, and DNA synthesis, by measuring the incorporation of 2H
into the respective end products for each pathway (119;128-132). For studies of lipid
2 metabolism, the use of this H2O methodology allows for an analysis of the integrated
flux of triglycerides over an extended time period in vivo, that yields a measure of tissue-
2 specific dynamics (119). Previous studies using H2O have revealed that there is considerable plasticity to the rates of triglyceride turnover in white adipose tissue in response to both nutritional and genetic factors (119;133-135).
Here we report that chronic ethanol feeding for 2-4 wks increased both the rates of triglyceride synthesis and degradation, but did not change the net deposition of
triglycerides in epididymal adipose (Table 3.2). Maintenance of net rates of triglyceride
- 54 - accumulation in epididymal adipose tissue after chronic ethanol feeding (Table 3.2) was
consistent with the equivalent rates of increase in epididymal fat content in ethanol- and
pair-fed rats (Fig 3.2). However, since the total adipose tissue weight was already
decreased after 2 weeks of ethanol feeding, these data suggest that there may have been a
decrease in the rate of fat accumulation during the first 2 weeks of ethanol feeding.
Triglyceride synthesis and degradation in adipose tissue are highly active and regulated processes; these processes modulate the net accumulation of lipid during growth. The fatty acids incorporated into adipose triglycerides originate either from de novo lipogenesis from carbohydrate or from the re-esterification of free fatty acids released during hydrolysis of triglycerides. In addition to the increased hydrolysis of triglycerides (Table 3.2), chronic ethanol feeding increases the rate of lipogenesis in rat adipose tissue (71). Lipogenesis in adipose tissue is stimulated by insulin (35;36), but inhibited by β-adrenergic agonists (136;137). Chronic ethanol desensitizes adipocytes to
β-adrenergic receptor activation (118) (Fig 3.5) and inhibits the sensitivity of adipocytes to insulin (59) (Fig 3.6). Therefore, increased lipogenesis in adipose tissue after chronic ethanol is likely due to an imbalance between β-adrenergic and insulin receptor-mediated responses. Taken together, these data suggest that both increased de novo lipogenesis (71) and increased availability of free fatty acids released during hydrolysis of triglycerides
(Table 3.2) may contribute to the increased rate of triglyceride synthesis after chronic ethanol feeding.
As with the hormonal regulation of lipogenesis, hydrolysis of triglycerides or lipolysis
in adipocytes is also regulated by hormones, initiated by the stimulation of β-adrenergic
receptors and inhibited by insulin (117). Since chronic ethanol feeding increased the rate
- 55 - of triglyceride degradation, we hypothesized that chronic ethanol either increased the
stimulatory response to β-adrenergic receptors and/or decreased the inhibitory response to
insulin. However, in isolated adipocytes, we have previously reported that β-adrenergic
receptor-stimulated lipolysis was actually suppressed, rather than enhanced, after chronic ethanol feeding (118). Therefore, based on the data from isolated adipocytes (118), we hypothesized that it was unlikely that increased β-adrenergic receptor activity after chronic ethanol contributed to increased lipolysis in vivo. This hypothesis was confirmed
in vivo by showing that β3-adrenergic receptor-stimulated systemic lipolysis was suppressed after chronic ethanol exposure (Fig 3.5).
Since chronic ethanol exposure did not increase β-adrenergic receptor-stimulated
lipolysis, we next investigated the effects of chronic ethanol on the anti-lipolytic action of
insulin using both ex vivo model of primary adipocytes, as well as an in vivo model utilizing a hyperinsulinemic-euglycemic clamp. Chronic ethanol feeding impaired the ability of insulin to inhibit lipolysis both ex vivo and in vivo. While hyperinsulinemic-
euglycemic clamps are commonly used to assess the sensitivity of glucose disposal to
insulin (125), this clamp technique is also used to estimate the sensitivity of systemic lipolysis to insulin using plasma glycerol Ra as an index (126). During the steady state of the clamp, plasma glycerol Ra was decreased by 28% from baseline in pair-fed rats, with no change in ethanol-fed rats (Fig 3.8). Consistent with the lack of decrease in glycerol
Ra, the ability of insulin to decrease plasma glycerol and free fatty acid concentrations
during the insulin clamp was also suppressed in ethanol-fed rats compared to pair-fed rats
(Fig 3.7).
- 56 - Chronic ethanol feeding impacts on a number of insulin-regulated metabolic pathways.
In addition to the loss of insulin-mediated inhibition of lipolysis after chronic ethanol
reported here, chronic ethanol feeding to rats also decreases whole-body glucose utilization during the hyperinsulinemic-euglycemic clamp (58;97), as well as insulin- stimulated glucose uptake in isolated rat adipocytes (59;98). In humans, short term
exposure to ethanol decreases peripheral glucose utilization, assessed either with
hyperinsulinemic-euglycemic clamps (91;92) or stable isotope analysis of gluconeogenic
flux during ethanol infusion (89). Studies in individual tissues and cell types have found
that ethanol impairs the insulin signaling pathway in a variety of cell types, including
cerebellar neurons (101), hepatocytes (102), and adipocytes (105), suggesting that chronic ethanol-induced insulin resistance likely results from impaired insulin signaling.
In addition to the reduced sensitivity to insulin observed after chronic ethanol feeding,
basal concentrations of plasma insulin were also decreased (Table 3.3). This decrease
may be due to an impaired function of pancreatic β cells by ethanol, as evidenced by the
inability of rats to maintain a second pulse of insulin release during a glucose tolerance
test after chronic exposure to ethanol (99;138;139). Taken together, these data suggest
that both the lower basal plasma insulin concentration, as well as the reduced ability of
insulin to inhibit lipolysis in adipocytes, contribute to the increased rate of triglyceride
degradation in adipose tissue after chronic ethanol feeding.
Because chronic ethanol feeding increased the rate of triglyceride degradation in
adipose tissue, it would also be expected that plasma free fatty acid concentrations would
be increased. However, we found no differences in the baseline concentration of plasma
free fatty acids between pair- and ethanol-fed rats (Fig 3.5B and Table 3.3). Maintenance
- 57 - of normal baseline concentrations of free fatty acids despite an increased rate of triglyceride degrdation suggests that the free fatty acids released during triglyceride degradation were either rapidly re-esterified in adipose tissue and/or rapidly taken up by other tissues once released into the circulation. This rate of uptake may actually be increased after chronic ethanol, as chronic ethanol consumption increases the hepatocellular uptake of long chain fatty acids (62). Interestingly, chronic ethanol feeding did increase plasma free fatty acid concentration during the steady state of the hyperinsulinemic-euglycemic clamp (Fig 3.7 C/D), suggesting that chronic ethanol may elevate free fatty acids in the circulation under certain conditions, such as hyperinsulinemia.
In summary, we have demonstrated that chronic ethanol feeding to rats increased the in vivo rates of triglyceride turnover in epididymal adipose tissue. Increased lipolytic capacity of adipose tissue after ethanol was not due to an increased sensitivity to β- adrenergic activation, but instead due to a suppression of the anti-lipolytic response of adipocytes to insulin. These data thus identify for the first time the mechanisms by which chronic ethanol consumption disrupts lipid homeostasis in adipose tissue. The dysregulation of adipose tissue metabolism thus likely contributes to the pathophysiologocal effects of ethanol.
- 58 - Chapter 4
Disruption of β-Adrenergic Receptor-Stimulated Lipolysis Pathway by Chronic
Ethanol
4.1. Introduction
Chronic ethanol exposure disrupts receptor-activated signal transduction in a variety
of cell types (48). One target of ethanol is the G protein-mediated signaling pathways
(48). The effect of ethanol on G protein-dependent responses is cell type specific.
Chronic administration of ethanol reduces the stimulation of cAMP concentration by
norepinephrine, and reduces the stimulation of adenynyl cyclase activity by the β-
adrenergic agonist isoproterenol in rodent brains (52;53). Conversely, in adipocytes,
ethanol feeding for 4 weeks causes a sensitization to stimulation by isoproterenol as well
as an increase in the quantity of immunoreactive Gαs protein (59).
Previous studies have demonstrated that chronic ethanol feeding decreases β3- adrenergic receptor agonist-stimulated systemic lipolysis, as measured by plasma free fatty acid and glycerol concentrations (Fig 3.5). However, the mechanism is unknown.
Since adipose tissue is the most active organ for lipolysis and lipolysis in adipocytes is regulated by β-adrenergic receptor signaling known to be affected by chronic ethanol, we hypothesized that β-adrenergic receptor regulation of lipolysis in adipose tissue may be susceptible to long-term ethanol exposure. In the present work, we report that four-week ethanol feeding to rats suppressed β-adrenergic receptor-mediated activation of lipolysis in adipocytes isolated from epididymal fat. The suppression of lipolysis in response to β-
- 59 - adrenergic activation was associated with a reduction of intracellular cAMP accumulation,
PKA activation, as well as phosphorylation of perilipin A and HSL.
4.2. Materials and Methods
4.2.1. Materials
Male Wistar rats (150-160g) were purchased from Harlan Sprague Dawley
(Indianapolis, IN). The Lieber-DeCarli high-fat ethanol diet was from Dyets (Bethlehem,
PA). Antibodies were obtained from the following sources: anti-PDE4 and anti-PDE4B from Abcam Inc. (Cambridge, MA), anti-phospho-(Ser/Thr) PKA substrate from Cell
Signaling Technology (Beverly, MA); anti-perilipin A/B from Research Diagnostics, Inc.
(Flanders, NJ); anti-caveolin from BD Transduction Laboratories (San Jose, CA); and
anti-ERK from Upstate (Charlottesville, VA). Maltose dextrins were purchased from
BioServ (Frenchtown, NJ), CompleteTM, EDTA-free protease inhibitor cocktail tablets
and enhanced chemiluminesence reagents from Roche (Indianapolis, IN), Calyculin A, cilostamide and Ro20-1724, a phosphodiesterase (PDE) 4 selective inhibitor, from
BIOMOL (Plymouth Meeting, PA). cAMP enzymeimmunoassay BiotrakTM (EIA)
system was from Amersham Pharmacia Biotech (Piscataway, NJ), PKA assay kit was
purchased from Upstate (Charlottesville, VA), [γ-32P]ATP was from PerkinElmer Life
Sciences (Boston, MA), [3H]cAMP was from Amersham Pharmacia Biotech (Piscataway,
NJ), and all other reagents were from Sigma (St. Louis, MO).
4.2.2. Animal care and feeding
- 60 - Rats were allowed ad libitum access to the Lieber-DeCarli ethanol diet or pair-fed a
control diet as previously described (98). Briefly, rats were housed in individual wire-
bottom cages under controlled temperature and humidity with 12 h light/dark cycle.
After arrival, rats were acclimatized for 3 days on rat chow and water. The rats were then
matched by weight and randomly assigned to the ethanol-fed or pair-fed groups, and
allowed free access to control liquid diet for two days (140). The ethanol-fed group had
ad libitum access to a liquid diet with 17% total calories as ethanol for 2 days and then
35% total calories as ethanol for 4 weeks. Controls were pair-fed a liquid diet that was
identical to the ethanol diet except that maltose dextrins were isocalorically substituted
for ethanol. Pair-fed rats were given the same amount of food as their ethanol-fed
counterparts consumed in the preceding 24 h. All procedures involving animals were
approved by the Institutional Animal Care and Use Committee at Case Western Reserve
University.
4.2.3. Isolation of adipocytes
After 4 weeks of feeding, rats were anesthetized by intraperitoneal injection of 0.075
mL for ethanol-fed rats or 0.12 mL for pair-fed rats per 100 g body weight of a cocktail
containing 10 mg/mL acepromazine, 100 mg/mL ketamine and 20 mg/mL xylazine. The
use of a lower dose of anesthetic for ethanol-fed rats was due to an increased sensitivity
of rats to the anesthetic cocktail after ethanol feeding. The effects of the use of different
anesthetic regimes for pair- and ethanol-fed rats on the lipolytic rate of adipocytes were
neglected since ketamine and xylazine have been shown to have either no effects or equivalent effects on lipolysis under the doses used in this study (121;122). Under
- 61 - anesthesia, rat epididymal fat pads were removed. Epididymal fat pads were selected as
the depot for this study because epididymal adipose tissue is more sensitive to
isoproterenol stimulation of lipolysis than subcutaneous fat (141). Adipocytes were then
isolated by collagenase digestion following the method of Rodbell (142) as modified by
Honner et al. (143), and as described before (59). Briefly, fat pads were weighed, washed
twice with Hanks’ balanced salt solution containing 25 mM N-2-hydroxyethylpiperazine-
N’-2-ethanesulfonic acid (HEPES), minced by scissors, and incubated at 37˚C in a
shaking water bath (80 revolutions/min (rpm)) for 50 min in 10 mL Hanks’ balanced salt
solution containing 10 mg collagenase (type II), 10 mg/mL bovine serum albumin (BSA)
(RIA grade), 25 mM HEPES, and 200 nM adenosine, pH 7.4. Adipocytes were then filtered through a 250 μm nylon mesh and washed twice with 15 mL phosphate-buffered saline (PBS) containing 1 mM sodium pyruvate and 200 nM adenosine, pH 7.4 (Wash buffer). Adipocytes were then counted, and cell concentrations adjusted to 1 x 106 cells/mL.
4.2.4. Ex vivo lipolysis assay and cAMP concentration
Adipocytes were isolated as described above except that adenosine was not added
during digestion and washing steps unless indicated in the figure legend. In addition, 1
mg/mL BSA (RIA grade) was included in Wash buffer. 200 μL aliquots of cells (1 x 106 cells/mL) were placed into 5 mL polypropylene tubes and a number of agents were added:
1) isoproterenol (10-10-10-5 M) with or without pre-incubation of cells with 10 µM Ro20-
1724 for 3 min; 2) adenosine deaminase (ADA) (0.4 U/mL) and (-)-N6-(2-
Phenylisopropyl)adenosine (R-PIA) (10 nM); 3) dibutyryl-cAMP (0.05-1 mM).
- 62 - Adipocytes were incubated for 1 h at 37˚C in a shaking water bath (100 rpm), and lipolysis was measured as glycerol release (144). After incubation, cells were centrifuged
briefly and the cell medium was collected for analysis of glycerol concentration using
glycerol reagent (GPO trinder reagent) in a flat-bottom 96-well plate. Optical density at
540 nm was measured on a Maxline microplate reader. Control experiments showed that
glycerol release from isolated adipocytes was dependent on incubation time (from 20 to
90 min) and cell concentration (from 0.5 to 1.8 x 106 cells/mL) in cells from both pair-
and ethanol-fed rats. For cAMP assay, 100 μL of cell suspension was removed to a 96-
well plate containing 10 μL of 2% NP40 in 1 N HCl after 0.5-15 min of incubation. The
samples were frozen at -20˚C until analysis of cAMP concentration by ELISA using the
cAMP enzymeimmunoassay BiotrakTM (EIA) system.
4.2.5. PDE activity
1 mL aliquots of isolated adipocytes were incubated with or without 1 µM
isoproterenol in the presence of 1 U/mL ADA for 5 or 10 min at 37˚C in a shaking water
bath (100 rpm) and then immediately placed on ice and homogenized in 500 μL ice cold
TSE buffer (20 mM Tris, 255 mM sucrose, 1 mM EDTA, 40 μL/mL protease inhibitor
cocktail, 10 μL/mL activated Na vanadate) with 60 nM calyculin A. Homogenates were
then briefly spun and the fluid fraction was collected. PDE activity was assayed by the
method of Ahmad et al. (145). Briefly, 20 μg samples were incubated with 0.1 μmol/L
[3H]cAMP for 10 min at 30˚C in the presence or absence of 1 μM cilostamide (PDE3B
inhibitor) or 10 μM Ro20-1724 (PDE4 inhibitor), and the formed [3H]AMP was degraded
to [3H]adenosine by 0.3 mg/mL 5’-nucleotidase of rattlesnake venom. [3H]adenosine
- 63 - was separated from the reaction mixture by chromatography on QAE sephadex A-25
columns, and the radioactivity in the eluate was measured by liquid scintillation counting.
PDE3B activity was expressed as the difference between activity with and without
cilostamide, and PDE4 activity was expressed as the difference between activity with and
without Ro20-1724.
4.2.6. PKA activity
PKA activity was measured using the Kemptide assay (146). 1 mL aliquots of
isolated adipocytes were treated with or without 1 µM isoproterenol for 2-10 min in the
presence of 1 U/mL ADA and cell homogenates were prepared in TSE buffer as
described above. The homogenates were then centrifuged at 5,500 g for 10 min at 4ºC and the fluid fraction collected. Protein content was measured and 20 µg protein was used for assessing PKA activity. The activity of PKA was measured over 10 min in the
presence of 0.33 µM PKC inhibitor peptide and 3.3 µM CaMK inhibitor peptide.
Maximal PKA activity was assessed by the addition of 1.67 µM cAMP.
4.2.7. Western blotting
Quantification of immunoreactive PDE4-1 mL aliquots of isolated adipocytes, treated with 1 U/mL ADA, were homogenized in the modified radioimmunoprecipitation (RIPA) buffer (50 mM Tris, pH 7.4, 1% NP-40, 150 mM NaCl, 1 mM EDTA, 40 μL/mL protease inhibitor cocktail, 1 mM Na3VO4, 1 mM NaF). After removal of the fat layer by
centrifugation at 4,000 g for 3 min, the fluid fraction was prepared in the SDS sample
buffer and applied to a 6% SDS-polyacrylamide gel. Immunoreactive PDE4 isoforms
- 64 - were probed using both PDE4 (detecting all known PDE4 A and D variants) and PDE4B
(detecting all known PDE4B proteins) antibodies.
Quantification of immunoreactive perilipin A/B and phospho-perilipin A and HSL-1 mL aliquots of isolated adipocytes were treated with or without 1 µM isoproterenol or 0.5 mM dibutyryl-cAMP for 5 or 10 min in the presence of 1 U/mL ADA and cell homogenates were prepared in TSE buffer. The homogenates were then centrifuged at
5,500 g for 10 min at 4ºC. After removal of the fluid fraction, 1 mL chloroform/methanol
(1:1) mixture was added to the fat layer. Samples were vortexed thoroughly and spun for
10 min at 16,000 g at 4ºC to precipitate fat layer-associated proteins. Protein pellets were then dissolved in 1% SDS and protein concentration was measured. Both cell homogenates and fat layer-associated protein samples were prepared in SDS sample buffer and applied to an 8% SDS-polyacrylamide gel. Proteins were then transferred to
PVDF membrane for Western blotting. Total perilipin A/B was probed using an anti- perilipin A/B antibody. Phospho-perilipin A and phospho-HSL were probed with a phospho-(Ser/Thr) PKA substrate antibody. Protein expression of ERK1 in adipocyte homogenates was not different between pair- (2.15 ± 0.3 arbitrary units of density, n = 4) and ethanol-fed rats (1.93 ± 0.3 arbitrary units of density, n = 4); therefore, ERK1 was used as a loading control of the cell homogenate samples (Fig 4.7/4.8). ERK1 was probed using an anti-ERK antibody. Caveolin is found in lipid droplets (147); therefore, we used caveolin as a control for equal protein extraction from fat samples. Caveolin was probed with anti-caveolin in a 12% duplicate gel of fat layer-associated protein samples.
- 65 - 4.2.8. Statistical analyses
Data are expressed as means ± SEM. Dose response curves of adipocytes to
isoproterenol were estimated by non-linear regression (GraphPad Prism® 4; San Diego,
CA). Statistical analyses were performed using the general linear model procedure on
SAS for personal computers. Differences between groups were determined by least square means.
4.3. Results
We have previously reported that chronic ethanol feeding increases Gαs expression associated with an increase in β-adrenergic receptor activation in adipocytes (59).
Therefore, we first asked if chronic ethanol feeding increased lipolysis in response to β-
adrenergic receptor activation. Basal rates of lipolysis did not differ between adipocytes
isolated from pair- (0.13 ± 0.02 μmol/106 cells; n = 8) and ethanol-fed (0.14 ± 0.02
μmol/106 cells; n = 8) rats. Lipolysis in adipocytes from both pair- and ethanol-fed rats
displayed a sigmoidal dose-response to isoproterenol (Fig 4.1A). EC50 values for
isoproterenol-stimulated lipolysis were not different between pair- (94.0 nM) and
ethanol-fed (81.0 nM) rats. However, maximal stimulation of lipolysis by isoproterenol
in adipocytes isolated from ethanol-fed rats was reduced to 29% of that observed in pair-
fed rats (Fig 4.1A). This attenuation in isoproterenol-stimulated lipolysis was not
influenced by extracellular adenosine since, in the presence of ADA (0.4 U/ml) and R-
PIA (an A1 receptor agonist, 10 nM), isoproterenol (1 µM)-stimulated lipolysis was still
lower in adipocytes from ethanol-fed rats compared to pair-fed rats (Fig 4.1B).
Additionally, reduced isoproterenol-stimulated lipolysis was not due to a reduced total
- 66 - A B
1.2 Pair-fed cells) cells) 6 * * 0.9
1.0 EtOH-fed 6 0.8 0.8 mol/10 0.7 mol/10 μ
* μ 0.6 0.6 0.4 0.5 0.4 0.2 Glycerol release 0.3 * 0.0 0.2 M)-stimulated glycerol release
-0.2 μ 0.1 -10 -9 -8 -7 -6 -5 (increase over basal, 10 10 10 10 10 10 0.0 Pair-fed EtOH-fed Isoproterenol (M) Iso (1 (increase over ADA/PIA,
Figure 4.1 Chronic ethanol feeding decreased β-adrenergic receptor-stimulated
lipolysis in adipocytes isolated from epididymal fat.
A) Adipocytes isolated from pair- and ethanol-fed rats were treated with or without
increasing concentrations of isoproterenol (Iso). Basal lipolysis, measured as glycerol
release over 1 hour, did not differ in adipocytes from pair- (0.13 ± 0.02 μmol/106 cells; n
= 8) and ethanol-fed (0.14 ± 0.02 μmol/106 cells; n = 8) rats. Values are the increase in glycerol release in response to isoproterenol over basal, and represent means ± SEM (n =
3), and were graphed using non-linear regression. B) Lipolysis in adipocytes was measured as described in “EXPERIMENTAL PROCEDURES” except that adenosine was included in the digestion and washing steps during adipocyte isolation. Adipocytes were treated with or without 1 µM isoproterenol in the presence of ADA (0.4 U/mL) and
R-PIA (an A1 receptor agonist, 10 nM). Values are the increase in glycerol release in response to isoproterenol over release in the presence of ADA and R-PIA, and represent means ± SEM (n = 4). *p < 0.05 Pair-fed vs. EtOH-fed.
- 67 - capacity of lipolysis in adipocytes, as maximal rate of lipolysis, measured in the presence
of ADA (148), did not differ between adipocytes from pair- (6.6 ± 0.9 µmol/106 cells; n =
4) and ethanol-fed (6.2 ± 0.7 µmol/106 cells; n = 4) rats.
Since cAMP is the key signal to increase HSL-dependent lipolysis, we asked whether
the suppression of β-adrenergic receptor-stimulated lipolysis after chronic ethanol
feeding was associated with decreased cAMP accumulation. Basal cAMP concentration
in adipocytes did not differ between pair- and ethanol-fed rats (Fig 4.2A). In response to
isoproterenol, cAMP concentration increased rapidly in adipocytes from pair-fed rats
with a peak at 1 min, followed by a rapid decline. In contrast, this peak of isoproterenol-
stimulated cAMP accumulation was suppressed after ethanol feeding. After stimulation
by isoproterenol for 5-15 min, cAMP concentrations remained elevated over basal, but
was not different between pair- and ethanol-fed rats (Fig 4.2A). cAMP accumulation is a
balance between its synthesis from ATP by activated adenylyl cyclase and its degradation
to AMP by activated phosphodiesterases (PDEs) (117). Chronic ethanol’s reduction in
the early peak of β-adrenergic receptor-dependent cAMP accumulation could be due to
either a decrease in cAMP synthesis or an increase in cAMP degradation. To measure
cAMP synthesis independent of degradation, adipocytes were pretreated with Ro20-1724,
a PDE4 selective inhibitor. In the presence of Ro20-1724, β-adrenergic receptor-
stimulated cAMP production was higher in adipocytes from both pair- and ethanol-fed
rats compared to the accumulation observed in the absence of Ro20-1724 (Fig 4.2B vs.
Fig 4.2A). In contrast to the inhibition of the early peak of cAMP in the absence of
Ro20-1724, cAMP production in the presence of Ro20-1724 was higher in ethanol-fed
rats after isoproterenol stimulation for 5-15 min compared to pair-fed rats (Fig 4.2B).
- 68 - A
40 * Pair-fed
ulation * EtOH-fed m 30 * * * cells) * 6 20 * * + + * (pmol/10 10 + * * ulated cAMP accu m
0 Iso-sti 0.0 2.5 5.0 7.5 10.0 12.5 15.0 Time (min)
B
Pair-fed 70 6 * EtOH-fed ulation 60 m * 50 * * 40 * * * 30 * * + 20 * + *
ulated cAMP accu * +
m 10
0
Iso-sti 0.0 2.5 5.0 7.5 10.0 12.5 15.0 in the presence of RO (pmol/10cells) Time (min)
Figure 4.2 Chronic ethanol feeding suppressed β-adrenergic receptor-stimulated
cAMP accumulation by increasing cAMP degradation via PDE4.
Adipocytes isolated from pair- and ethanol-fed rats were pre-treated with (B) or without
(A) 10 μM Ro20-1724 (Ro) for 3 min, then 1 μM isoproterenol was added except the basal samples and incubation was continued for another 0.5-15 minutes. Intracellular
cAMP concentration was determined by cAMP enzymeimmunoassay BiotrakTM system.
Data are means ± SEM (n = 3~7). *p < 0.05 compared to 0 min within dietary group; +p
< 0.05 compared to pair-fed.
- 69 - These data suggest that chronic ethanol feeding decreased cAMP accumulation primarily through increased cAMP degradation via PDE4, rather than decreased cAMP synthesis.
To further clarify the involvement of PDE4 in chronic ethanol-reduced cAMP accumulation, PDE activity was measured. Adipocytes predominantly express two PDE isoforms, PDE3B and PDE4 (149;150). Basal activity of PDE4 was significantly higher after chronic ethanol feeding compared to pair feeding (Fig 4.3A). Isoproterenol increased PDE4 activity in cell homogenates from pair-fed rats, but did not further increase PDE4 activity in cell homogenates from ethanol-fed rats (Fig 4.3A). In contrast to the increase in basal activity of PDE4, chronic ethanol feeding had no effect on either basal or isoproterenol-stimulated activation of PDE3B (Fig 4.3B).
PDE4 is a multi-gene family. Four genes (PDE4A, PDE4B, PDE4C, and PDE4D) encode over 18 different PDE4 isoforms (151). Since chronic ethanol feeding increased the activity of PDE4, we asked whether this increase was due to the up-regulation of expression of one or more of PDE4 family members. Western blot analysis of PDE4 using a PDE4 antibody (detecting all known PDE4 A and D variants) and a PDE4B antibody (detecting all known PDE4B variants), showed that the quantity of immunoreactive PDE4A, PDE4B, or PDE4D isoforms was not increased by chronic ethanol; indeed, the expression of some isoforms was even decreased in adipocytes from ethanol-fed rats compared to pair-fed rats (Fig 4.4).
Since chronic ethanol feeding decreased β-adrenergic receptor-stimulated intracellular cAMP accumulation, we next asked whether the activity of PKA, a downstream target of cAMP, was also impaired by chronic ethanol feeding. Basal activity of PKA was not affected by chronic ethanol feeding (Fig 4.5A/B). Isoproterenol dose-dependently
- 70 -
A Pair-fed EtOH-fed b 7
g) b
m 6 in/
m 5 b ol/
m 4
3
2 a
1 PDE4 activity (p 0 Basal Isoproterenol
B Pair-fed EtOH-fed 20 g) m in/
m 15 ol/ m 10
5
PDE3B activity (p 0 Basal Isoproterenol
Figure 4.3 Chronic ethanol feeding increased basal activity of PDE4.
Adipocytes isolated from pair- and ethanol-fed rats were incubated with or without 1 µM isoproterenol for 5 min. Cell homogenates were used to analyze PDE activity as described in “EXPERIMENTAL PROCEDURES”. Data represent means ± SEM (n = 8 for basal samples or 4 for isoproterenol-treated samples). Values with different letters are significantly different (p < 0.05).
- 71 - A Rat testes Pair-fed EtOH-fed
100 KDa
75 KDa
B Rat testes Pair-fed EtOH-fed
100 KDa
75 KDa
Figure 4.4 Chronic ethanol feeding did not increase the quantity of immunoreactive
PDE4A, PDE4B, or PDE4D isoforms.
Adipocytes isolated from pair- and ethanol-fed rats were homogenized in RIPA buffer.
After removal of the fat layer by centrifugation at 4,000 g for 3 min, the fluid fraction was prepared in SDS sample buffer and applied to a 6% SDS-polyacrylamide gel. PDE4 was immunoblotted using a PDE4 antibody (detecting all known PDE4 A and D variants)
(A) and a PDE4B antibody (detecting all known PDE4B variants) (B). Rat testes were used as positive controls. A representative immunoblot from 6 experiments is shown.
Asterisks indicate the immunoreactive PDE4 isoforms detected in rat testes, and the arrowheads indicate the corresponding bands detected in rat adipocytes.
- 72 - A Pair-fed * 5000 EtOH-fed
4000
3000 PKA activity d 2000 * 1000 + + ( pmol/min/mg) Iso-stimulate 0 0.0 2.5 5.0 7.5 10.0 Time (min) B 17500 Pair-fed
g) EtOH-fed
m 15000 in/
m 12500 ol/
m 10000
7500
5000
2500 PKA activity (p 0 No cAM P 1.67μMcAMP
Figure 4.5 Chronic ethanol feeding decreased β-adrenergic receptor-stimulated
PKA activity, but not maximal PKA activity.
Adipocytes isolated from pair- and ethanol-fed rats were incubated with (A) or without (B)
1 µM isoproterenol for 0-10 min. Cell homogenates were used to analyze PKA activity as described in “EXPERIMENTAL PROCEDURES”. Maximal PKA activity was assessed by the addition of 1.67 µM cAMP (B). Data represent means ± SEM (n =3~8).
*p < 0.05 compared to own basal; +p < 0.05 Pair-fed vs. EtOH-fed.
- 73 - increased PKA activity in cell homogenates from pair-fed rats. However, PKA activation
in response to isoproterenol was inhibited by chronic ethanol feeding (Fig 4.5A). In the
presence of 1.67 µM cAMP, maximal activation of PKA did not differ between pair-
feeding and ethanol-feeding (Fig 4.5B).
Upon activation, PKA phosphorylates multiple substrates in adipocytes, but two
particular substrates have emerged as important in the regulation of lipolysis, perilipin A
and HSL (117). Since chronic ethanol feeding decreased isoproterenol-stimulated PKA
activity, we next investigated whether isoproterenol-stimulated phosphorylation of
perilipin A and/or HSL was also decreased by ethanol. To further investigate the
subcellular distribution of these proteins, we isolated the fat layer from cell homogenates
and extracted fat layer-associated proteins. Using caveolin, a protein resident in the lipid
droplet as a marker, we found equivalent recovery of lipid droplet protein from
adipocytes from both pair- and ethanol-fed rats (Fig 4.7). Adipocytes express two forms
of perilipin, A and B. Western blot analysis showed that both forms of perilipin were
predominantly localized to the fat layer and their expression was not affected by chronic
ethanol feeding (Perilipin A: 2.42 ± 0.53 arbitrary units of density in pair-fed, 2.13 ± 0.47
in ethanol-fed; perilipin B: 0.40 ± 0.15 arbitrary units of density in pair-fed, 0.35 ± 0.14
in ethanol-fed; n = 3).
To assess the phosphorylation of perilipin A and HSL, western blot analysis with a
phospho-(Ser/Thr) PKA substrate antibody was utilized. Phosphorylation of a 62 KDa protein, which migrated coincident with immunoreactive perilipin A, was identified as phospho-perilipin A (Fig 4.6A). Phosphorylation of an 83 KDa protein, the exclusive protein in the molecular weight range of 75 to 200 KDa detected by phospho-(Ser/Thr)
- 74 - A Iso (min) 0 5 10 0 5 10 KDa Peri A 50 50 p-Ser/Thr Peri B 37 37
B Iso (min) 0 5 10 KDa: 200
150
100 p-Ser/Thr 75
Figure 4.6 Phosphorylation of perilipin A and HSL were detected by a phospho-
(Ser/Thr) PKA substrate antibody.
Adipocytes isolated from pair-fed rats were treated with or without 1 μM isoproterenol
for 5 or 10 min, and fat layer-associated protein samples or whole cell homogenates were
prepared as indicated. A) Fat layer-associated protein samples were subjected to SDS-
PAGE and immunoblotted with antibodies against perilipin A/B (Peri A/B) or phospho-
(Ser/Thr) PKA substrate (p-Ser/Thr). B) Cell homogenates were subjected to SDS-PAGE and immunoblotted with phospho-(Ser/Thr) PKA substrate antibody.
- 75 - A Fat layer Homogenate Figure 4.7 Chronic ethanol PF EF PF EF Iso (min) 0 5 10 0 5 10 0 5 10 0 5 10 feeding reduced β-adrenergic p-Peri A p-Peri A
Caveolin ERK 1 receptor-stimulated phosphorylation of perilipin A 5 Pair-fed EtOH-fed 4 and HSL. * Adipocytes isolated from pair- 3 * 2 and ethanol-fed rats were treated
phospho-Perilipin A * 1 with or without 1 μM (arbitrary units of density) 0 Iso (min) 0 5 10 0 5 10 isoproterenol for 5 or 10 min, and Fat layer Homogenate fat layer-associated protein B Fat layer Homogenate PF EF PF EF samples or whole cell Iso (min) 0 5 10 0 5 10 0 5 10 0 5 10 p-HSL p-HSL homogenates were prepared as
Caveolin ERK 1 indicated. Phospho-perilipin A Pair-fed 2.0 EtOH-fed (p-Peri A) (A) and phospho-HSL
1.5 (p-HSL) (B) were immunoblotted
* using a phospho-(Ser/Thr) PKA 1.0 * substrate antibody. A) A phospho-HSL 0.5 * representative immunoblot is (arbitrary units of density) 0.0 Iso (min) 0 5 10 0 5 10 shown with longer exposure for Fat layer Homogenate homogenate compared to fat layer. With equal exposure, phospho-perilipin A was 3.8 arbitrary units of density in fat layer compared to 1 in homogenate. B) A representative immunoblot is shown with equal exposure. Caveolin was used as a control for equal protein extraction from fat samples;
ERK1 was used as a loading control in the cell homogenates. Values represent means ±
SEM (n = 4~6). *p < 0.05 Pair-fed vs. EtOH-fed. PF, pair-fed; EF, ethanol-fed.
- 76 - PKA substrate antibody in adipocyte homogenates, which had the same molecular weight
as HSL, was identified as phospho-HSL (Fig 4.6B). Phospho-perilipin A was not
detectable at baseline in either the fat layer or cell homogenates from pair- and ethanol-
fed rats (Fig 4.7A). However, treatment of adipocytes from pair- and ethanol-fed rats with isoproterenol for 5 or 10 min induced a strong and rapid phosphorylation of perilipin
A; this response was more pronounced in the fat layer (Fig 4.7A). Concomitant with
decreased isoproterenol-stimulated PKA activation, phosphorylation of perilipin A was
inhibited by chronic ethanol feeding in the fat layer (Fig 4.7A). Similarly,
phosphorylated HSL was not detectable in non-stimulated cells, but was induced in
response to isoproterenol (Fig 4.7B). Phosphorylation of HSL was decreased in both the
fat layer and cell homogenates from ethanol-fed rats compared to pair-fed rats after stimulation with isoproterenol. In contrast to the abundance of phospho-perilipin A in the
fat layer, phospho-HSL was distributed in both the fat layer and non-fat fraction (Fig
4.7B).
Decreased β-adrenergic receptor-stimulated phosphorylation of perilipin A and HSL
after ethanol feeding could result from decreased cAMP accumulation or a combination
of decreased cAMP accumulation and a direct impact of ethanol on protein
phosphorylation. To ask whether ethanol directly impaired phosphorylation of these
proteins, independent of changes in cAMP concentration, we determined phosphorylation
of perilipin A and HSL in the presence of dibutyryl-cAMP, a PDE-resistant cAMP analog.
Treatment of adipocytes with dibutyryl-cAMP for 5 or 10 min stimulated
phosphorylation of both perilipin A and HSL (Fig 4.8). In contrast to the inhibition of
isoproterenol-stimulated phosphorylation of perilipin A and HSL after chronic ethanol
- 77 - A Fat layer Homogenate PF EF PF EF db-cAMP (min) 0 5 10 0 5 10 0 5 10 0 5 10 p-Peri A p-Peri A
Caveolin ERK 1
B Fat layer Homogenate PF EF PF EF db-cAMP (min) 0 5 10 0 5 10 0 5 10 0 5 10 p-HSL p-HSL
Caveolin ERK 1
Figure 4.8 Chronic ethanol feeding did not affect dibutyryl-cAMP-stimulated phosphorylation of perilipin A or HSL.
Adipocytes isolated from pair- and ethanol-fed rats were treated with or without 0.5 mM
dibutyryl-cAMP (db-cAMP) for 5 or 10 min, and fat layer-associated protein samples or
whole cell homogenates were prepared as indicated. Phospho-perilipin A (p-Peri A) (A)
and phospho-HSL (p-HSL) (B) were immunoblotted using a phospho-(Ser/Thr) PKA
substrate antibody. A representative immunoblot from 4 experiments is shown. PF, pair-
fed; EF, ethanol-fed.
- 78 - feeding, dibutyryl-cAMP-stimulated phosphorylation of perilipin A and HSL did not
differ between pair- and ethanol-fed rats in either the fat layer or cell homogenates (Fig
4.8).
Since treatment of adipocytes with dibutyryl-cAMP restored ethanol-reduced
phosphorylation of proteins, we then asked whether suppressed lipolysis could also be
restored by the addition of dibutyryl-cAMP. Treatment of adipocytes with dibutyryl-
cAMP increased lipolysis in adipocytes isolated from both pair- and ethanol-fed rats in a
dose-dependent manner (Fig 4.9A). However, dibutyryl-cAMP-stimulated lipolysis in
adipocytes from ethanol-fed rats was consistently lower than that in adipocytes from pair-
fed rats at all doses tested (Fig 4.9A). Furthermore, since decreased cAMP accumulation
after ethanol was restored in the presence of PDE4 inhibitor, Ro20-1724 (Fig 4.2B), we
examined whether the decrease in isoproterenol-stimulated lipolysis after ethanol feeding
was also restored when PDE4 activity was inhibited. While pretreatment of adipocytes
with Ro20-1724 did not change isoproterenol-stimulated lipolysis in adipocytes from
pair-fed rats, it elevated lipolysis in adipocytes from ethanol-fed rats. However,
adipocyte lipolysis in ethanol-fed rats was still lower compared to pair-fed rats in the
presence of Ro20-1724 (Fig 4.9B).
4.4. Discussion
Adipose tissue is the major site for storage of triglycerides; mobilization of free fatty
acids and glycerol via lipolysis provides a rapid source of fuel for other organs in response to fasting, infection and inflammation (33). During chronic ethanol exposure,
whole-body lipid homeostasis is disrupted, with the eventual development of hepatic
- 79 - A Pair-fed B Pair-fed EtOH-fed EtOH-fed 1.4 * 1.00 d d cells) 6 1.2 * cells) 6 0.75 1.0 c mol/10 μ 0.8 mol/10 μ 0.50 b 0.6 * + 0.4 + + 0.25 0.2 a a 0.0 Glycerol release ( 0.00 0.00 0.05 0.10 0.50 1.00 Glycerol release ( Basal Iso Iso + Ro db-cAMP (mM)
Figure 4.9 Chronic ethanol feeding impaired signaling downstream of phosphorylation of perilipin A and HSL.
Adipocytes isolated from pair- and ethanol-fed rats were treated with or without increasing concentrations of dibutyryl-cAMP (db-cAMP) (A) or pretreated with or without 10 μM Ro20-1724 (Ro) for 3 min prior to stimulation with 1 μM isoproterenol
(Iso) (B). Lipolysis was determined as glycerol release over 1 hour. Values represent means ± SEM (n =3~8). *p < 0.05 compared to own basal; +p < 0.05 Pair-fed vs. EtOH- fed. Values with different letters are significantly different (p < 0.05).
- 80 - steatosis (60). While it is clear that the regulation of lipolysis in adipocytes plays an
important role in maintaining lipid homeostasis, the influence of chronic ethanol feeding
on the regulation of lipolysis in adipocytes has not been investigated. Here we report that
chronic ethanol feeding to rats decreased β-adrenergic receptor-stimulated lipolysis. This
suppression of β-adrenergic activation of lipolysis was associated with inhibition of
intracellular cAMP accumulation and PKA activation, as well as decreased
phosphorylation of perilipin A and HSL. cAMP signaling is known to be an important
target of acute and chronic ethanol exposure in a number of systems (48). In adipocytes,
chronic ethanol feeding impaired β-adrenergic signaling at least two sites; first, chronic
ethanol feeding increased PDE4 activity, leading to decreased cAMP accumulation and
second, impaired PKA-mediated phosphorylation of perilipin A and HSL, two proteins
localized to the lipid droplet of adipocytes. These data thus are the first to identify an
association between impaired PKA-mediated phosphorylation of proteins at the lipid
droplet with decreased lipolysis during a pathophysiological response to diet, such as
chronic ethanol feeding. Taken together, these data provide evidence that chronic ethanol
feeding results in a complex dysregulation of β-adrenergic regulation of lipolysis and suggest that impaired regulation of lipolysis in adipose tissue may contribute to the pathophysiological effects of ethanol on lipid homeostasis.
Chronic ethanol feeding to rats had no effect on basal rates of lipolysis in adipocytes isolated from epididymal adipose tissue, but diminished β-adrenergic receptor-stimulated lipolysis compared to pair feeding (Fig 4.1). These data are consistent with a previously reported decrease in adrenaline-induced lipolysis in adipocytes from rats provided ethanol in their drinking water for 2 weeks (72). Inhibition of β-adrenergic receptor-
- 81 - stimulated lipolysis after chronic ethanol feeding was due, at least in part, to decreased accumulation of intracellular cAMP (Fig 4.2A). Chronic ethanol-induced desensitization of β-adrenergic receptor-cAMP signaling has been reported in a number of other cell types, primarily associated with changes in expression of heterotrimeric G proteins (48).
In adipocytes, we have previously reported that chronic ethanol feeding increases
expression of Gαs, and this increase is associated with an increase in β-adrenergic
receptor-stimulated cAMP synthesis, when measured in the presence of a PDE4 selective inhibitor (Fig 4.2B) (59). However, despite these increased rates of cAMP production, accumulation of cAMP by adipocytes after chronic ethanol feeding in response to β- adrenergic activation was actually decreased (Fig 4.2A), suggesting an increase in cAMP degradation via PDE4 after chronic ethanol feeding. Direct measurement of PDE activity showed increased activity of PDE4 in adipocytes from ethanol-fed rats, with no effect of ethanol feeding on activity of PDE3B (Fig 4.3). This increase in PDE4 activity was not due to increased expression of PDE4A, PDE4B, or PDE4D (Fig 4.4). Of the 11 known
isoforms of PDE, PDE4 and PDE3 are the predominant isoforms expressed in the
adipocytes (149). However, these two isoforms are differentially localized and regulated
within rat adipocytes. PDE4, originally found in the cytosolic fraction and recently
shown to interact with β-arrestin and thus be recruited to plasma membrane, is regulated
by PKA- and MAPK-mediated phosphorylation (47), while PDE3, localized in the
microsomal fraction, is activated by insulin (46). In this study, PDE4 activity was
specifically increased after chronic ethanol exposure, functionally countering the ethanol-
induced increase in Gαs/cAMP synthesis, and decreasing cAMP accumulation. While
one other report has described an increase in PDE4 activity by acute ethanol in bovine
- 82 - bronchial epithelial cells (152), this is the first report of a change in PDE activity after
chronic ethanol feeding and suggests that there is a pathophysiological relationship
between the regulation of PDE4 and the impact of ethanol on cellular processes.
In addition to the decrease in isoproterenol-stimulated cAMP accumulation, signaling
downstream of cAMP, PKA activation and PKA-dependent phosphorylation of perilipin
A and HSL in response to isoproterenol, were also decreased by chronic ethanol feeding
(Fig 4.5/4.7). Perilipin is a lipid droplet-associated protein in the PAT protein family,
including perilipin, adipocyte differentiation-related protein (ADRP), and TIP47 (153).
These structurally and functionally similar PAT proteins play a critical role in lipid
metabolism in a variety of cell types (154-156). Two forms of perilipin are expressed in
rat adipose tissue, perilipin A and B. Perilipin A, the dominant isoform in adipocytes not
only acts as a protective barrier in the absence of stimulation, but is necessary to facilitate
HSL translocation from the cytosol to the surface of lipid droplet when it is
phosphorylated by PKA (157). By contrast, perilipin B fails to protect basal lipolysis or
elicit an activation response (158). In the present study, total quantity of perilipin A and
B was not affected by chronic ethanol feeding; however, phosphorylation of perilipin A
in response to isoproterenol was abolished in adipocytes from ethanol-fed rats compared
to pair-fed rats (Fig 4.7A). Upon lipolytic stimulation of adipocytes, a redistribution of
perilipin from the surface of lipid droplet to the cytosol has been reported (159;160).
This redistribution of perilipin may be subject to regulation. Clifford et al. (159) found no movement of perilipin from the lipid droplet to the cytosol in isoproterenol-stimulated adipocytes isolated from young rats (180-220g); however, there was a significant movement of perilipin away from the lipid droplet in adipocytes isolated from more
- 83 - mature rats (230-280g). In contrast, in our rat model using mature rats (280-340g), both unphosphorylated and phosphorylated forms of perilipin were predominantly localized to the fat layer of adipocytes (Fig 4.7A and data not shown). Thus, chronic ethanol feeding impaired the phosphorylation of perilipin A, rather than its distribution within the adipocyte.
Phosphorylation of perilipin A and HSL in response to isoproterenol was decreased
after chronic ethanol feeding; however, phosphorylation of these proteins per se was not
affected by ethanol since dibutyryl-cAMP-stimulated phosphorylation of perilipin A and
HSL was not different between pair-feeding and ethanol-feeding (Fig 4.8). Despite
normal phosphorylation of perilipin A and HSL in response to dibutyryl-cAMP, lipolysis
in adipocytes treated with dibutyryl-cAMP was still decreased by chronic ethanol feeding
(Fig 4.9A). Additionally, lipolysis in adipocytes pretreated with a PDE4 inhibitor, Ro20-
1724, was lower in ethanol-fed rats than pair-fed rats (Fig 4.9B). These results suggest
that, in addition to reduced cAMP accumulation and cAMP-dependent PKA activation and phosphorylation of perilipin A and HSL, chronic ethanol feeding impaired the
signaling pathway from phosphorylation of perilipin A and HSL to the eventual stimulation of lipolysis. Upon phosphorylation, HSL translocates from the cytosol to the
surface of lipid droplets, initiating the hydrolysis of triglycerides (38). While the target
of ethanol is not known, impaired translocation of HSL from the cytosol to the surface of
lipid droplets is a possible site of action.
In summary, the current investigations demonstrate that chronic ethanol feeding to rats
decreased β-adrenergic receptor-stimulated lipolysis in adipocytes isolated from
epididymal adipose tissue. This reduction was associated with inhibited intracellular
- 84 - cAMP accumulation and coincident repression of cAMP-dependent PKA activation and phosphorylation of perilipin A and HSL. Further, decreased β-adrenergic receptor- stimulated cAMP accumulation was attributable to increased PDE4 activity after chronic ethanol feeding. These data suggest that the disruption of β-adrenergic receptor regulation of lipolysis may contribute to changes in whole-body lipid homeostasis seen after chronic ethanol exposure.
- 85 - Chapter 5
Disruption of Glucose Disposal by Chronic Ethanol
5.1. Introduction
Epidemiological studies have shown that patients with type 2 diabetes have disrupted glucose homeostasis, as evidenced by postprandial and fasting hyperglycemia, impaired glucose tolerance test, as well as impaired insulin-stimulated glucose utilization during a hyperinsulinemic-euglycemic clamp (161). Chronic heavy alcohol consumption, as an independent risk factor for type 2 diabetes, also disrupts glucose homeostasis, associated with the development of insulin resistance (59;97;98). Chronic ethanol consumption in both human and rodents decreases whole-body glucose utilization during a hyperinsulinemic-euglycemic clamp (58;96;97). These data suggest that the disruption of glucose homeostasis may provide a specific mechanism by which chronic heavy ethanol increases the risk for type 2 diabetes.
While adipose tissue and skeletal muscle are the two major sites of glucose utilization in response to insulin, the effects of ethanol on the glucose transport in these particular tissues are not completely understood. Previous studies demonstrated that chronic ethanol feeding to rats decreases insulin-stimulated glucose uptake in isolated adipocytes
(59;98), but not in isolated skeletal muscle (99). However, the in vivo effects of chronic ethanol on insulin’s action on glucose transport have not been studied extensively. While one study reported that two weeks of ethanol feeding to rats impaired glucose utilization and increased hepatic glucose production during the hyperinsulinemic-euglycemic clamp, the effects of chronic ethanol on tissue-specific glucose disposal were not investigated
- 86 - (97). In the present work, we utilized hyperinsulinemic-euglycemic clamps, coupled with the use of isotopic tracers, to determine the relative contributions of skeletal muscle and adipose tissue in mediating impaired glucose disposal after chronic ethanol exposure.
5.2. Materials and Methods
5.2.1. Materials
Male Wistar rats (150-160g) were purchased from Harlan Sprague Dawley
(Indianapolis, IN). The Lieber-DeCarli high-fat ethanol diet was purchased from Dyets
(Bethlehem, PA). Maltose dextrins were obtained from BioServ (Frenchtown, NJ).
13 [ C6]glucose (99 atom percent excess) were purchased from Isotec (Miamisburg, OH), bis(trimethylsilyl)trifluoroacetamide + 10% trimethylchlorosilane was purchased from
Regis Technologies Inc. (Morton Grove, IL), gas chromatography-mass spectrometry
(GC-MS) supplies were from Agilent Technologies (Wilmington, DE). Blood glucose meter and blood glucose test strips were purchased from CVS (Woonsocket, RI), human insulin was from Eli Lilly (Indianapolis, IN), rat insulin ELISA was from Mercodia Inc.
(Winston Salem, NC), and all other reagents were from Sigma (St. Louis, MO).
5.2.2. Animal care and feeding
Male Wistar rats were allowed free access to the Lieber-DeCarli high-fat ethanol diet
or pair-fed a control diet for 4 weeks as previously described (118). Randomly assigned
ethanol-fed rats were provided an ad libitum liquid diet containing ethanol as 36% of
total calories for 4 weeks. Control rats were pair-fed a liquid diet that was identical to the
ethanol diet except that maltose dextrins were isocalorically substituted for ethanol. Pair-
- 87 - fed rats were given the same amount of food as their ethanol-fed counterparts consumed
in the preceding 24 h. All procedures involving animals were approved by the
Institutional Animal Care and Use Committee at Case Western Reserve University.
5.2.3. Hyperinsulinemic-euglycemic clamp
After 3 wks of pair- or ethanol-feeding, rats were anesthetized by inhalation of an
isoflurane and oxygen mixture and the left carotid artery and the right jugular vein were
catheterized for blood sampling and intravenous infusion during the clamp, respectively.
All rat surgeries were done in the Mouse Metabolic and Phenotyping Center at Case
Western Reserve University. Rats were allowed a week to recover from the surgery
while maintained on their respective diets. Hyperinsulinemic-euglycemic clamps were
performed on one rat at a time as previously described (125), with minor modifications.
Rats were transported from the Animal Resource Center and allowed at least 90 min to
13 stabilize before commencement of the glucose clamp. [ C6]Glucose (~1 µmol/kg/min)
was continuously infused from the jugular vein catheter for 90 min basal period and 2-h
clamp period. Baseline levels of blood glucose and plasma insulin were determined as
the mean of values obtained in blood samples collected at -30 and -5 min. At time 0, a
primed (60 mU/kg)/continuous (4 mU/kg/min) infusion of human insulin was started and
continued for 2 hrs and 45 mins. The blood glucose concentration was clamped at euglycemic level by a variable rate infusion of 20% glucose. Blood glucose was monitored with a blood glucose meter and the rate of glucose infusion adjusted every 10 min. Blood samples for determination of plasma insulin were obtained at -30, -5, 10, 20,
30, 40, 50, 60, 70, 80, 90, 100, 110, and 120 min, and blood samples for determination of
- 88 - 13 plasma [ C6]glucose enrichment were obtained at -30, -5, 90, 100, 110, and 120 min. At
120 min, 50 µCi of 2-deoxy-D-[1, 2-3H]glucose ([3H]2DG) was administrated as an
intravenous bolus. Blood samples were collected at 2, 5, 10, 15, 30, and 45 min after the
bolus administration. After the last blood sampling, rats were anesthetized and the
epididymal adipose tissue, subcutaneous adipose tissue, omental adipose tissue, soleus
muscle, and gastrocnemius muscle were removed and frozen in liquid nitrogen.
5.3.4. Glucose utilization rate
The rate of plasma glucose turnover was calculated by the following equation: Ra =
(ENRinf/ENRpl-1) · F, where ENRinf is the isotopic enrichment of the infusate, ENRpl is
the isotopic enrichment of plasma and F is the rate of the isotope infusion (126). The
13C-labeling of plasma glucose was determined as described below. 20 µl of plasma was
deproteinized with 200 µl of methanol by centrifugation for 10 min at 16,100 x g. The
fluid fraction was then evaporated to dryness and reacted with 50 µl hydroxylamine (25
mg hydroxylamine in 1 ml of pyridine) for 20 min at 75°C, followed by a reaction with
50 µl of bis(trimethylsilyl)trifluoroacetamide + 10% trimethylchlorosilane for 20 min at
75°C. Isotope enrichment was determined by GC-MS under electron impact ionization;
ions of mass-to-charge ratios (m/z) 319-323 were monitored.
5.2.5. Radioactivity measurements of plasma and tissue samples
Plasma and tissue samples were processed as previously described (162). Briefly, 10
µl of plasma was deproteinized with 100 µl of barium hydroxide (0.1 N) and 100 µl of zinc sulfate (0.1 N) in the presence of 10 µl of saline. The mixture was vortexed and
- 89 - centrifuged at 2,300 x g for 5 min. The radioactivity of the supernatant was then determined by liquid scintillation counting. Tissue samples were weighed, homogenized in 0.5% perchloric acid, and centrifuged at 2,300 x g for 5 min, and the supernatants were neutralized with KOH. One aliquot of homogenate was counted without further treatment to yield total counts of [3H]2DG and 2-deoxy-D-[1, 2-3H]glucose phosphate
([3H]2DG-P). A second aliquot of homogenate was treated with barium hydroxide (0.3 N) and zinc sulfate (0.3 N) to remove [3H]2DG-P, and then was counted to yield [3H]2DG radioactivity. The tissue radioactivity of [3H]2DG-P was calculated by the difference of
total counts ([3H]2DG and [3H]2DG-P) and the [3H]2DG count alone.
5.2.6. Uptake of 2-deoxy-[3H]glucose in isolated adipocytes
After 4 weeks of feeding, rats were anesthetized and adipose depots including epididymal, subcutaneous, and omental adipose tissue were removed. Adipocytes were then isolated by collagenase digestion as previously described (59), counted and diluted
6 to 1x10 cells/ml in phosphate-buffered saline with 1 mM MgCl2, 0.68 mM CaCl2, 1
mg/ml BSA, 1 mM pyruvic acid, and 1 U/ml adenosine deaminase, pH 7.4. Adipocytes
were stimulated with or without 10 nM insulin for 30 min at 37°C, and uptake of
[3H]2DG (final concentration of 2DG 2.5 mM, 0.33 µCi/tube) was measured for 3 min
(59). Nonspecific uptake was measured in the presence of 0.3 mM phloretin.
5.2.7. Statistical analyses
- 90 - Data are expressed as mean ± SEM. Statistical analyses were performed using either
student’s t test or the general linear model procedure on SAS for personal computers.
Differences between groups were determined by least square means.
5.3. Results
Since the disruption of glucose homeostasis has been proposed as a specific
mechanism by which chronic ethanol consumption increases the risk for type 2 diabetes,
here we used the hyperinsulinemic-euglycemic clamp technique to determine the effects
of chronic ethanol on glucose homeostasis in vivo. Body weights of rats used in the
hyperinsulinemic-euglycemic clamps did not differ between pair- (275 ± 5 g, n = 8) and
ethanol-fed (256 ± 10 g, n = 7) rats. There was no difference between pair- and ethanol-
fed rats in baseline level of blood glucose (Fig 5.1A). Blood glucose concentration was
maintained at euglycemia during 90-120 min of the clamp and did not differ between
pair- and ethanol-fed rats (Fig 5.1A). Baseline concentration of plasma insulin was lower
in ethanol-fed rats compared to pair-fed rats, possibly due to an impaired insulin secretion
from β-cells after ethanol exposure (99;138;139). The primed/continuous infusion of
insulin increased plasma insulin concentration and achieved steady state at 90-120 min of
the clamp in both pair- and ethanol-fed rats; the steady state level of plasma insulin was not different between pair- and ethanol-fed rats (Fig 5.1B).
In order to maintain euglycemia during the hyperinsulinemic-euglycemic clamp, 20%
glucose was continuously infused at a variable rate (Fig 5.2A). Plasma insulin was
maintained at steady state (Fig 5.1B) and glucose infusion rate was also constant (Fig
5.2A) in both pair- and ethanol-fed rats during the 90-120 min of the clamp. Therefore,
- 91 - A 200 Pair-fed EtOH-fed
g/dl) 150 m
100
50 Blood glucose (
0 0 90 100 110 120 Time of the clamp (min) B 1000 Pair-fed EtOH-fed
ol/l) 750 m
500 a insulin (p
m 250 Plas * 0 * 0 20 40 60 80 100 120 Time of the clamp (min)
Figure 5.1 Rat blood glucose and plasma insulin levels during hyperinsulinemic-
euglycemic clamps.
A) The blood glucose concentration was clamped at euglycemic level by a variable rate
infusion of 20% glucose, and was monitored with a blood glucose meter. B) Human insulin was primed-continuously infused for 2 hours. Plasma insulin level was determined by ELISA. Data represent means ± SEM (n = 8 for pair-fed group and 7 for ethanol-fed group). *p < 0.05 Pair-fed vs. Ethanol-fed.
- 92 - A B C
in) 30 Pair-fed 30 30 m EtOH-fed g/kg/ m 20 20 20 * * , mg/kg/min)
10 90-120 10 10 (GIR Glucose infusion rate Glucose utilization rate
0
0 (mean at 90-120 min,0 mg/kg/min) Glucose infusion rate ( 0 20 40 60 80 100 120 Pair-fed EtOH-fed Pair-fed EtOH-fed Time of the clamp (min)
DE 8 * 90 7 80 6 70 60 5 * 50 4 , mg/kg/min) 40 3 90-120 30 the clamp (%) 2 20 (HGP 1 10 Hepatic glucose production Suppression of HGP during 0 0 Pair-fed EtOH-fed Pair-fed EtOH-fed
Figure 5.2 Chronic ethanol feeding decreased glucose infusion rate, glucose
utilization rate, and percent suppression of hepatic glucose production during the
hyperinsulinemic-euglycemic clamp.
A) To maintain euglycemia, 20% glucose was infused at a variable rate during the clamps.
B) Mean glucose infusion rate (GIR90-120) was presented as the mean of 90, 100, 110, and
120 min values as the steady state values. C) Glucose utilization rate at the steady state of the clamp was presented as the mean of 90, 100, 110, and 120 min values. D) Hepatic glucose production (HGP) was determined as the difference between glucose utilization rate and glucose infusion rate. HGP90-120 represents the mean of 90, 100, 110, and 120
min values. E) The percent suppression of hepatic glucose production during the clamp
was calculated. Data represent means ± SEM (n = 8 for pair-fed group and 7 for ethanol-
fed group). *p < 0.05 Pair-fed vs. EtOH-fed.
- 93 - the last 30 min of the clamp was assumed to be at steady state and the means of the 90,
100, 110, and 120 min values were used for steady-state values (126). The mean glucose
infusion rate at the steady state was lower in ethanol-fed rats compared to pair-fed rats
(Fig 5.2B).
The baseline level of plasma glucose turnover rate did not differ between pair- and
ethanol-fed rats (data not shown). However, at the steady state of the clamp, plasma
glucose turnover/utilization rate was lower in ethanol-fed rats compared to pair-fed rats
(Fig 5.2C), suggesting that chronic ethanol feeding impaired the whole-body glucose
utilization in response to insulin. Subtracting the glucose infusion rate from the glucose
turnover rate, hepatic glucose production at the steady state of the clamp was calculated.
Ethanol-fed rats had higher hepatic glucose production compared to pair-fed rats (Fig
5.2D). Therefore, the percent suppression of hepatic glucose production by insulin was
impaired by chronic ethanol feeding (Fig 5.2E). The decreased glucose utilization rate
combined with impaired suppression of hepatic glucose production during the
hyperinsulinemic-euglycemic clamp suggests that chronic ethanol feeding induced
insulin resistance.
Skeletal muscle and adipose tissue are the two major sites of glucose disposal in
response to insulin. To elucidate the contribution of different tissues mediating impaired
whole-body glucose disposal, we determined tissue-specific glucose disposal using a
non-metabolizable glucose analog, [3H]2DG. The bolus administration of [3H]2DG
maximally increased plasma [3H]2DG at 2 min, then the radioactivity of plasma [3H]2DG declined over time in both pair- and ethanol-fed rats (Fig 5.3A). However, the radioactivity of plasma [3H]2DG was not different between pair-and ethanol-fed rats (Fig
- 94 - A BC Pair-fed Pair-fed Pair-fed 600 120 EtOH-fed EtOH-fed 1200 EtOH-fed 500 100 900 400
l) 80 μ 300 600 60 (dpm/ H]glucose phosphate H]glucose in plasma
3 200 3 H]glucose phosphate
* (dpm/mg tissue) 3 40 *
300 (dpm/mg tissue) * 100 20 2-deoxy-[ 2-deoxy-[ 0 0 2-deoxy-[ Soleus Gastro 0 10 20 30 40 50 0 Epi SubQ Oment Time of the clamp (min)
Figure 5.3 Chronic ethanol feeding decreased glucose disposal in adipose tissue, but
not in skeletal muscle during the hyperinsulinemic-euglycemic clamp.
A) At 120 min of the clamp, [3H]2DG was administrated as an intravenous bolus. The
radioactivity of [3H]2DG in plasma was determined by liquid scintillation counting from
samples obtained at 2, 5, 10, 15, 30, and 45 min after the bolus. B, C) After the last blood
sampling, rats were anesthetized and epididymal adipose tissue (Epi), subcutaneous
adipose tissue (SubQ), omental adipose tissue (Oment), soleus muscle (Soleus), and
gastrocnemius muscle (Gastro) were collected. The radioactivity of [3H]2DG-P in
individual tissues was then measured. Data represent means ± SEM (n = 9 for pair-fed
rats and 3 for ethanol-fed rats). *p < 0.05 Pair-fed vs. EtOH-fed.
- 95 - 5.3A). Over 45 min, [3H]2DG was transported, phosphorylated and accumulated in
tissues. The radioactivity of [3H]2DG-P was decreased in epididymal, subcutaneous, and
omental adipose tissues in rats after ethanol feeding compared with pair feeding (Fig
5.3B). In contrast, the [3H]2DG-P radioactivity in skeletal muscle, either soleus or gastrocnemius muscles, was not affected by ethanol feeding (Fig 5.3C).
Consistent with the data obtained from the clamps, insulin-stimulated glucose
transport in isolated adipocytes was also decreased by chronic ethanol feeding (Fig 5.4)
(59;98). Insulin increased glucose transport in adipocytes isolated from epididymal,
subcutaneous, and omental adipose tissues in pair-fed rats, but did not stimulate transport
in adipocytes from ethanol-fed rats (Fig 5.4). Taken together, these data suggest that
chronic ethanol feeding decreased whole-body glucose disposal by impairing the
utilization of glucose by adipose tissue.
5.4. Discussion
Chronic ethanol consumption disrupts glucose homeostasis in both human and animal
models, as evidenced by decreasing whole-body glucose utilization during
hyperinsulinemic-euglycemic clamps (58;97) (Fig 5.2C). While adipose tissue and
skeletal muscle are the two major organs utilizing glucose in response to insulin, the
relative contribution of these two tissues to impaired glucose utilization after chronic
ethanol has not been investigated. Here we reported that chronic ethanol feeding to rats
decreased glucose utilization during the hyperinsulinemic-euglycemic clamp, and this
- 96 - 5000 Pair-fed EtOH-fed 4000
cells) 3000 5
H]glucose uptake 2000 3
(pmol/10 1000 * * * 0 2-deoxy-[ BIns BIns BIns
Epi SubQ Oment
Figure 5.4 Chronic ethanol feeding decreased insulin-stimulated glucose uptake in adipocytes isolated from epididymal, subcutaneous, and omental adipose tissues.
Adipocytes isolated from epididymal (Epi), subcutaneous (SubQ), and omental (Oment) adipose tissues in rats after 4 weeks of pair- or ethanol-feeding, were treated with (Ins) or without (B) 10 nM insulin for 30 min. Uptake of 2-deoxy-[3H]glucose was then measured. Data represent means ± SEM (n = 4 for epididymal adipose tissue, 5 for subcutaneous adipose tissue, and 3 for omental adipose tissue). *p < 0.05 Pair-fed vs.
EtOH-fed.
- 97 - decrease was associated with impaired glucose transport in adipose tissues, but not in skeletal muscle. These results for the first time identify the importance of adipose tissue, rather than skeletal muscle, in the pathogenesis of diet-induced insulin resistance in response to chronic ethanol feeding.
Four-week ethanol feeding to rats decreased the glucose appearance rate, an indicator of glucose utilization rate, at the steady state of the hyperinsulinemic-euglycemic clamp
(Fig 5.2C). These data are consistent with previous reports of decreased whole-body glucose utilization in response to insulin in rats either chronically fed with ethanol- containing liquid diet or intragastrically administrated with ethanol (58;97), suggesting that chronic consumption of ethanol induces systemic insulin resistance. This impaired insulin sensitivity by ethanol has also been observed in humans. Hyperinsulinemic- euglycemic clamp studies involving chronic heavy drinkers or alcoholics have demonstrated decreased peripheral glucose utilization compared to non-drinking controls
(96). Chronic ethanol-induced insulin resistance was also characterized by an impairment of the ability of insulin to inhibit hepatic glucose production during the hyperinsulinemic-euglycemic clamp (Fig 5.2 D/E). Impaired suppression of hepatic glucose production by insulin is consistent with elevated hepatic gluconeogenesis measured in isolated liver slices from male rats after chronic ethanol consumption (163).
Since adipose tissue and skeletal muscle are the two major organs utilizing glucose in response to insulin, we further determined the relative contribution of these two tissues to impaired glucose utilization after chronic ethanol. During the hyperinsulinemic- euglycemic clamp, chronic ethanol feeding decreased glucose disposal in adipose tissue including epididymal, subcutaneous, and omental adipose depots. However, the glucose
- 98 - disposal in two different types of skeletal muscles, soleus and gastrocnemius muscles was
not affected by chronic ethanol (Fig 5.3). These data are also consistent with the ex vivo studies of glucose transport in isolated adipocytes or muscle cells. Chronic ethanol exposure to rats decreases insulin-stimulated glucose uptake in isolated adipocytes
(59;98;105) (Fig 5.4), but not in isolated epitrochlearis muscle (99). Taken together,
these results suggest that chronic ethanol feeding decreases whole-body glucose
utilization during the hyperinsulinemic-euglycemic clamp by impairing the glucose
utilization in adipose tissue, but not in skeletal muscle.
Different adipose depots exhibit varied metabolic properties and respond differentially
to nutrient and hormones upon specific metabolic demands in the body (164-166). For
example, increased lipolytic responses to catecholamines and decreased insulin-mediated
inhibition of lipolysis in visceral fat cells, compared to subcutaneous fat cells, causes
elevation of the free fatty acid levels in the portal blood and is associated with increased
risk to develop cardiovascular complications in the male type of obesity (167). We find
that both visceral and subcutaneous adipose depots exhibit a similar response to chronic
ethanol consumption. Chronic ethanol feeding decreases the expression and secretion of
adiponectin, an anti-inflammatory adipokine, in both subcutaneous and visceral adipose
tissues (168). Here we report that insulin-stimulated glucose transport was globally decreased in subcutaneous fat, omental fat (a visceral fat depot), as well as epididymal fat
(a specialized fat depot that partially exhibits the metabolic characteristics of both
subcutaneous and visceral fat), by chronic ethanol feeding (Fig 5.3B and Fig 5.4).
Under normal physiological conditions, adipose tissue takes up only a small fraction
of glucose compared to muscle (169) (Fig 5.3 B/C), which may explain the unchanged
- 99 - decay of [3H]2DG in plasma of pair- and ethanol-fed rats (Fig 5.3A). Although adipose
tissue contributes relatively little to total body glucose disposal, adipose-specific GLUT4
knockout mice develop a secondary insulin resistance in muscle and liver (170),
suggesting the presence of circulating factors that are released from adipose tissue
leading to insulin resistance.
In contrast to the adipose-specific GLUT4 knockout mouse model, chronic ethanol
feeding induced insulin resistance in adipose tissue, but insulin sensitivity in skeletal
muscle was preserved. While the mechanisms by which chronic ethanol feeding decreased insulin-stimulated glucose uptake in adipose tissue are not completely understood, studies of the regulation of insulin-mediated glucose transport in isolated adipocytes reveal that chronic ethanol feeding decreases the expression of GLUT4, the glucose transporter isoform that is expressed exclusively in adipocytes and muscle cells and is responsible for insulin-stimulated glucose transport (59). Insulin-stimulated fusion of GLUT4 vesicles to the plasma membrane is also disrupted by chronic ethanol feeding
(98). Although insulin signaling through PI 3-kinase is not a target of chronic ethanol
(98), the decrease in insulin-stimulated glucose transport by chronic ethanol is associated with disrupted insulin-mediated Cbl/TC10 signaling pathway in adipocytes (105). Thus, a combination of decreased GLUT4 expression and impaired GLUT4 trafficking likely leads to decreased glucose uptake in response to insulin in adipose tissue.
Despite the difference in insulin sensitivity of skeletal muscle, both adipose-specific
knockout of GLUT4 and chronic ethanol consumption exhibit either depleted or
decreased GLUT4 expression in adipose tissue and insulin resistance in adipose tissue,
suggesting that there might be similar mechanisms for the pathogenesis of insulin
- 100 - resistance under these two conditions. It is proposed that the presence of circulating
factors released from adipose tissue contributes to insulin resistance in adipose-specific
GLUT4 knockout mouse model (170). Therefore, it is also possible that changes in circulating concentrations of cytokines and adipokines, including increased TNFα, that are produced either from macrophages or adipose tissue, along with decreased adiponectin secretion from adipocytes, may contribute to the development of insulin resistance after chronic ethanol exposure (168). In addition to TNFα and adiponectin, several other adipose-derived proteins have been identified and found to be associated with obesity-induced insulin resistance, such as macrophage chemoattractant protein 1 and retinol-binding protein 4 (171-173). The effect of chronic ethanol feeding on expression of these adipose-derived molecules is currently under investigation.
In summary, here we have demonstrated that chronic ethanol feeding to rats decreased
whole-body glucose utilization and impaired the suppression of hepatic glucose
production by insulin during a hyperinsulinemic-euglycemic clamp. This decrease in
insulin-stimulated glucose utilization after chronic ethanol was associated with impaired
glucose transport in adipose tissue, rather than skeletal muscle. These data indicate that
chronic ethanol feeding disrupts glucose homeostasis in adipose tissue, likely
contributing to the pathophysiological effects of ethanol, such as type 2 diabetes.
- 101 - Chapter 6
Overall Summary and Future Prospects
6.1. Overall Summary
Chronic ethanol consumption induces hepatic steatosis and is a known risk factor for type 2 diabetes (9). While the mechanisms by which chronic ethanol triggers the progression of these pathophysiological conditions are unknown, the disruption of lipid and glucose homeostasis during ethanol exposure likely represents a potential contributor
(8). Because adipose tissue acts as a regulator for maintaining whole-body lipid and glucose homeostasis, we hypothesized that lipid and glucose homeostasis in adipose tissue might be vulnerable to chronic ethanol exposure. The current study has demonstrated that chronic ethanol exposure to rats disrupts both lipid and glucose
homeostasis in adipose tissue (Fig 6.1), likely providing a specific mechanism of the
pathology of ethanol.
Chronic ethanol consumption in rats increases the turnover rate of triglyceride in
epididymal adipose tissue, including both the rate of triglyceride synthesis and
degradation. The turnover of triglycerides in adipose tissue regulates both the lipid and energy homeostasis of whole body. This process is tightly controlled by a number of hormones including catecholamines and insulin. While the effects of chronic ethanol on the hormonal regulation of triglyceride synthesis have not been investigated, chronic ethanol feeding decreases β-adrenergic receptor-stimulated lipolysis both in vivo and in adipocytes isolated from epididymal adipose tissue. This decrease in the β-adrenergic receptor-stimulated lipolysis after chronic ethanol is associated with inhibition of
- 102 - ADIPOSE TISSUE PLASMA LIVER
TG Glycerol GNG HGP Hepatic ↑L ipo Glucose insulin lys ? is FFA resistance Adipose ↑ ↑DNL insulin resistance FFA ↓Oxidation ↑TG ↓VLDL
Systemic Insulin Resistance
Figure 6.1 The current model of chronic ethanol’s effects on lipid and glucose
homeostasis. Chronic ethanol exposure increases the rate of triglyceride degradation, which releases excess glycerol and free fatty acids into the blood stream. Increased availability of free fatty acids in the circulation combined with increased rate of de novo lipogenesis in liver, decreased rate of fatty acid oxidation in liver, and decreased secretion of very low density lipoprotein from the liver leads to accumulation of lipid in the liver or steatosis. Furthermore, chronic ethanol feeding induces adipose- and hepatic- insulin resistance, which along with elevated plasma free fatty acid concentration contributes to systemic insulin resistance seen after chronic ethanol feeding. ↑ increase; ↓ decrease; TG, triglyceride; FFA, free fatty acid; GNG, gluconeogenesis; HGP, hepatic glucose production; DNL, de novo lipogenesis; VLDL, very low density lipoprotein.
- 103 - intracellular cAMP accumulation and PKA activation, as well as decreased
phosphorylation of perilipin A and HSL. Instead, chronic ethanol feeding suppresses the
ability of insulin to inhibit lipolysis both in vivo utilizing hyperinsulinemic-euglycemic
clamps in conscious rats and ex vivo using isolated adipocytes. These results suggest that chronic ethanol feeding increases the lipolytic capacity of adipose tissue by suppressing the anti-lipolytic response to insulin, but not by decreasing the sensitivity of adipocytes to
β-adrenergic activation.
The increased rate of triglyceride degradation in adipose tissue after chronic ethanol
feeding likely contributes to the elevated plasma free fatty acid concentration (70), as
well as increased flux of fatty acids into the liver seen after chronic ethanol (62). The
combination of increased flux of fatty acids into the liver along with additional ethanol-
induced perturbations in the metabolic handling of fatty acids by the liver, including
increased rate of de novo lipogenesis (71), increased esterification of fatty acids to form
triglycerides and phospholipids (63), decreased oxidation of fatty acids (63), and
decreased secretion of very low density lipoprotein (65), contributes to the accumulation
of lipid in the liver or steatosis after chronic ethanol feeding.
In addition to the disruption of lipid metabolism, chronic ethanol consumption also
disrupts glucose metabolism in adipose tissue. Chronic ethanol feeding to rats decreases
whole-body glucose utilization during a hyperinsulinemic-euglycemic clamp. This
decreased glucose utilization after chronic ethanol mediates impaired glucose uptake in
adipose tissue, but not in skeletal muscle. Taken together, the data from these studies
demonstrate that chronic ethanol feeding to rats for 4 weeks induces adipose-insulin
resistance, as evidenced by impaired ability of insulin to inhibit lipolysis and stimulate
- 104 - glucose uptake in adipose tissue. Moreover, chronic ethanol feeding also impairs the
ability of insulin to inhibit hepatic glucose production. The insulin resistance in both
adipose tissue and liver leads to the systemic insulin resistance seen after chronic ethanol
consumption.
6.2. Future Prospects
While the current study raises many interesting questions for future research, one of
the most interesting questions to pursue is to investigate the mechanisms by which
chronic ethanol consumption induces insulin resistance. Chronic ethanol feeding induces
systemic insulin resistance, associated with insulin resistance in adipose tissue and liver,
but not in skeletal muscle. However, the mechanisms are still unclear. Additionally,
although adipose tissue is one of the major organs utilizing glucose as fuel in response to
insulin, it only takes up a small fraction of glucose compared to skeletal muscle under
normal physiological conditions (169). Therefore, it is not clear how the impaired
glucose disposal in adipose tissue alone, without decreased glucose disposal in skeletal muscle, is sufficient to impair whole-body glucose utilization.
The disruption of insulin signaling is a fundamental mechanism in the development of
insulin resistance. Chronic ethanol feeding, however, does not impair PI-3 kinase-
dependent insulin signaling prior to PI-3 kinase activation in liver, skeletal muscle, or
adipose tissue (97;98). Recent study in our lab indicated that chronic ethanol feeding
disrupts Cbl/TC10-mediated insulin signaling in adipocytes (105), another insulin-
mediated signaling pathway that has been shown in some studies to be required for
insulin-stimulated GLUT4 translocation and glucose uptake (83;84). Therefore,
- 105 - investigations of the effects of chronic ethanol feeding on insulin signaling downstream
from PI-3 kinase and Cbl/TC10-medited insulin signaling in various tissues might be
importance of understanding the mechanisms of chronic ethanol-induced insulin
resistance.
Abel et al. recently reported that adipose-specific GLUT4 knockout mice develop
insulin resistance secondarily in muscle and liver (170), associated with an increase in
circulating retinol binding protein 4 (RBP4), a putative “glucose sensing” peptide
secreted from adipose tissue (172). Chronic ethanol feeding also decreases GLUT4 expression in adipocytes by 35% and induces insulin resistance in adipose tissue (59).
However, skeletal muscle does not develop insulin resistance (99); perhaps because the
decrease in GLUT4 expression in muscle after chronic ethanol is modest. These data suggest that there may be similar mechanisms for the pathogenesis of adipose-specific
GLUT4 knockout- and chronic ethanol-induced insulin resistance. One potential
common mechanism may be related to changes in the expression of specific regulatory
molecules secreted by adipocytes.
Adipose tissue, which was originally considered to be an inert tissue functioning
solely as an energy store, is emerging as a key component of the immune system. A
variety of genes for secretory proteins are expressed in adipose tissue. Those include
various factors of the complement pathway, cytokines, chemokines, growth factors and
vasoactive substances such as plasminogen activator inhibitor in the fibrinolytic system
(174). These adipose tissue-derived bioactive substances have well-known roles in the
immune response. Importantly, several cytokines, such as tumor-necrosis factor α (TNFα)
and interleukin-6 (IL-6), and chemokines, such as macrophage chemoattractant protein 1
- 106 - (MCP-1) also have central roles in the regulation of insulin sensitivity, as well as many
respects of inflammation (174). Increased circulating concentrations, as well as increased
expression, of TNFα (175;176), IL-6 (177;178), and MCP-1 (171;179) in adipose tissue
is associated with insulin resistance in diet-induced and genetic models of obesity.
Retinol binding protein 4 (RBP4) is a recently identified adipose-derived molecule that is
also associated with insulin resistance, obesity, and type 2 diabetes (172;173). The serum concentration of RBP4 is increased in insulin-resistance states and elevated RBP4 levels
cause insulin resistance (172). In contrast to the increase of TNFα, IL6, MCP-1, and
RBP4, expression of adiponectin, an anti-inflammatory adipokine in adipose tissue, as well as its serum concentration is decreased in obesity and insulin resistance (180-183).
Chronic ethanol consumption is known to disrupt the expression and/or circulating
concentrations of some of these adipose-derived substances. For example, chronic ethanol feeding to rats increases the expression of TNFα mRNA in adipose tissue (168).
Circulating TNFα is also increased in the blood of alcoholics and animals chronically
exposed to ethanol (168;184;185). In addition to TNFα, chronic ethanol in humans and
rodents also increases other cytokines in the circulation, including IL-1β and IL-6 (175).
Conversely, expression of adiponectin in adipose tissue and its serum concentration are
decreased in rats chronically exposed to ethanol (168). These data suggest that chronic
ethanol consumption disrupts the profile of adipose-derived factors, which might act as
signals for insulin resistance. Therefore, further understanding of the effects of chronic
ethanol on these adipose-derived molecules might be of importance in understanding the
pathogenesis of ethanol-induced insulin resistance. Normalizing the concentration of
- 107 - adipose-derived particular proteins could be a new strategy for treating ethanol-induced insulin resistance and possibly other ethanol-related diseases.
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- 124 - Appendix 1
Protein Expression of PDE3B at Crude Plasma Membrane
Pair-fed EtOH-fed Ins (min) 0 1 2 5 10 0 1 2 5 10
PDE3B
Pair-fed
2.5 EtOH-fed
2.0
1.5
1.0
0.5
Protein expression of PDE3B 0.0 0 1 2 5 10 Stimulation of insulin (min)
Figure A1 The expression of PDE3B protein at the crude plasma membrane.
Adipocytes isolated from pair- and ethanol-fed rats were homogenized in TSE buffer, and the homogenates were spun at 15, 000 x g for 15 min to bring down the plasma membrane-enriched fraction at 4ºC. The pellets were then resuspended in 50 µl of TSE buffer, prepared in SDS sample buffer, and applied to SDS-polyacrylamide gel. Proteins were then detected by Western blotting. Ins, insulin. Data represent means ± SEM (n =
3-4).
- 125 - Appendix 2
Insulin-Stimulated Activation of PDE3B
b 30 b g) Pair-fed m 25 EtOH-fed in/ m
ol/ 20 a a m 15
10
5
PDE3B activity (p 0 Basal Insulin
Figure A2 Neither basal nor insulin-stimulated activity of PDE3B was affected by chronic ethanol feeding.
Adipocytes isolated from pair- and ethanol-fed rats were incubated with or without 10 nM insulin for 10 min. Twenty micrograms of cell homogenates were used to analyze
PDE3B activity as previously described. Data represent means ± SEM (n = 4). Values with different letters are significantly different (p < 0.05).
- 126 -