THE EXPRESSION AND ANTILIPOLYTIC ROLE OF
PHOSPHODIESTERASE 4 IN RAT ADIPOCYTES IN VITRO
DISSERTATION
Presented in Partial Fulfillment of the Requirement for
the Degree Doctor of Philosophy in the Graduate
School of The Ohio State University
By
Hong Wang, M.S.
* * * * *
The Ohio State University
2005
Dissertation Committee: Approved by
Dr. Mark Failla, Advisor
Dr. Neilé Edens Advisor Dr. Joshua Bomser The Ohio State University Dr. Martha Belury Nutrition Graduate Program
ABSTRACT
Elevated concentrations of plasma free fatty acids (FFA) may cause insulin
resistance. Inhibition of lipolysis reduces FFA availability and improves insulin
sensitivity. Lipolysis is stimulated by increased concentration of cAMP.
Phosphodiesterases (PDEs) hydrolyze cAMP and limit stimulation of lipolysis.
Phosphodiesterase 3B (PDE3B) is the major isoform of PDE in rat adipocytes and
mediates the antilipolytic effect of insulin. Phosphodiesterases 4 (PDE4) activity has also been detected in rat adipocytes, but its expression and physiological role are not clear.
In the first component of this dissertation research, the antilipolytic effect of Korean
ginseng extract (Panax ginseng; KGE) in rat adipocytes and the signaling pathway for
KGE antilipolysis were investigated. Adipocytes were isolated from rat adipose tissue by
collagenase digestion. The ability of KGE to inhibit lipolysis was assessed by measuring
glycerol release into the incubation medium. PDE inhibitors were applied to investigate
the signaling pathway for KGE antilipolysis. The present study showed that insulin
inhibited lipolysis by 42.4% compared to the basal lipolysis (p<0.002). The specific
PDE3 inhibitor cilostamide completely reversed insulin antilipolysis, whereas the specific
PDE4 inhibitor rolipram did not reduce insulin antilipolysis. KGE inhibited lipolysis by
49% compared to the basal lipolysis (p<0.002). Cilostamide and rolipram reduced KGE antilipolysis to 43% and 23.4% compared to basal conditions with the inhibitors,
ii respectively. Moreover, combination of the PDE3 and PDE4 inhibitors completely reversed KGE antilipolysis. In contrast with insulin, KGE did not affect phosphorylation of protein kinase B (PKB). These data suggest that ginseng mimicked the antilipolytic
effect of insulin through an alternative signaling pathway. KGE also partially affected
PDE4-mediated antilipolysis in rat adipocytes.
In the second component of this project, the expression of PDE4 in rat adipocytes
was investigated at the gene and protein levels. Reverse transcription-polymerase chain
reaction (RT-PCR) using published primers allowed amplification of fragments encoding
PDE3B (530bp), PDE4A (233bp), PDE4B (786bp), PDE4C (539bp), and PDE4D
(262bp) sequences. Real-time quantitative reverse transcription polymerase chain
reaction (QPCR) using the designed primers and probes to a region of high similarity
among PDE3B and PDE4 (A, B, C, D) sequences demonstrated that the expression of
four PDE4 subtypes (A, B, C, D) relative to PDE3B was 7%, 19%, 19%, and 7%,
respectively. PDE4B protein may be present in the cytosolic fraction of adipocytes. To
our knowledge, this is the first report that all four PDE4 subtypes are expressed in rat
adipocytes.
The final component of this project addressed the role of PDE4 in antilipolysis
6 mediated by prostaglandin E2 (PGE2) and N -(2-phenylisopropyl)adenosine (PIA). PGE2
inhibited lipolysis by 84% compared to the basal condition (p<0.006). Cilostamide and
rolipram reduced PGE2 antilipolysis to 76.3% and 46.6% compared to basal conditions
with the inhibitors, respectively. The combination of cilostamide and rolipram reduced
PGE2 antilipolysis to 17.5 %. PIA inhibited lipolysis by 92.2% compared to the basal lipolysis (p<0.006). Although cilostamide and rolipram alone did not affect PIA
iii antilipolysis, combination of these two inhibitors reduced PIA antilipolysis to 56.8%.
Moreover, the combination of two inhibitors has a synergistic effect on both PGE2 antilipolysis and PIA antilipolysis. In addition, cilostamide and rolipram increased basal lipolysis by 28% and 33.2%, respectively.
Subsequently, the results of PDE activity assay showed that isoproterenol, insulin, and PGE2 increased the total PDE activity in the particulate fraction of rat adipocytes by
65.2% (p<0.01), 30.6% (p<0.01), and 15.7% (p<0.01), respectively. In contrast, PIA
decreased the total PDE activity by 17.1% (p<0.01) and KGE did not change the total
PDE activity. PGE2 increased PDE3 activity by 21.4% (p<0.01), and PDE4 activity by
26.7% (p>0.3). These results suggest that PDE4 played a tonic role in antilipolysis, and
PGE2 antilipolysis was mediated in part by activation of PDE4 in rat adipocytes.
In summary, all four PDE4 subtypes were expressed in rat adipocytes. PDE4 partially mediated ginseng and PGE2 antilipolysis in rat adipocytes. Ginseng mimicked
the antilipolytic effect of insulin through an alternative signaling pathway. The
mechanism by which ginseng inhibits lipolysis was similar to PGE2 antilipolysis.
iv
DEDICATION
This dissertation is dedicated to my family for their love, support, and encouragement through the years.
v
ACKNOWLEDGMENTS
I would like to express my sincere gratitude to my two advisors, Dr. Neilé Edens and
Dr. Mark Failla, for their mentoring and support over the years. Dr. Edens’s dedication,
guidance, encouragement, and patience enabled my successful completion of this
dissertation. Dr. Failla’s intellectual challenge and insightful advice enlighten my
thinking, and his passion for work inspires me.
I would also like to express my deep appreciation to my committee members: Dr.
Joshua Bomser and Dr. Martha Belury for giving me valuable advice on my dissertation
research, and sharing their time and expertise with me over the years.
I would like to thank the scientists at Ross: Lisa Reaves for teaching me adipocyte
isolation and lipolysis assay, Irene Reyzer for helping me set up PDE experiment at Ross,
Mary-Beth Skeldling for teaching me glucose tolerance test on rats. I am very grateful to my best friend, Dr. Xincheng Zheng, for his technical assistance on RT-PCR. I would like to extend my appreciation to the scientists at Abbott, Dr. Paul Jung, Sandra Hu, and Dr.
Paul Kroeger, for their technical assistance on QPCR. I gratefully acknowledge Dr.
Manganiello and his two postdoctoral fellows, Dr. Faiyaz Ahmad and Dr. Young-Hun
Choi, at NIH for providing me the PDE3B antibody, and giving me critical assistance on
PDE assay.
vi I would like to thank many faculty and staff in the Department of Human Nutrition for their attention to my dissertation research. I am particularly grateful to Jessica
Pritchard for her handling my lab orders and facilitating my progress to graduation. I am
very grateful to Dr. Tammy Bray and her team for my training in her lab in the beginning
two years. I would like to thank all present and former graduate students in our
department, particularly Jordan Li, Chureeporn Chitchumroonchokchai, Li-Fen Liu, and
Xingya Wang, for their assistance and friendship all these years. I would like to express
my special gratitude to Dr. Shounan Yao for his careful proofreading of this dissertation.
I would like to especially thank my husband and my son for their love and consistent
support. My husband has given me a voluntary transportation between Ross and OSU for
my experiments the past two years, and shared much time and energy with our son. My
son, Weijia, can well understand his mother and does everything great in his school. Last,
I want to give my special thanks to my parents for their love, support, and expectation. I
will tell them that my Ph. D degree is for them.
vii
VITA
June 28, 1967…………………Born in Beijing, P. R. China
1986-1992…………………… Bachelor of Medicine, Beijing University of Chinese Medicine
1992-2000…………………… Physician, Department of Internal Medicine, China-Japan Friendship Hospital
1996-1999…………………… M.S., Beijing University of Chinese Medicine
2000-2005……………………Graduate Research Associate, The Ohio State University Graduate Nutrition Program
PUBLICATIONS
1. Xiaolin Tong, Hong Wang, Liying Cao, Fengling Wei, Fang Wan, and Zhiwen Ye. Effect of Yangyin injection on animal model of endotoxic fever. China Journal of Traditional Chinese Medicine and Pharmacy. 1998, 13(5): 30-33. (In Chinese)
2. Hong Wang, Fang Wan, Xiaolin Tong, and Zhiwen Ye. Effect of Yangyin injection on cell injury caused by endotoxin. China Journal of Traditional Chinese Medicine and Pharmacy. 2000, 15(1): 24-26. (In Chinese)
3. Jun Li, Hong Wang, Gary D. Stoner, and Tammy M. Bray. Dietary supplementation with cysteine prodrugs selectively restores tissue glutathione levels and redox status in protein-malnourished mice. Journal of Nutritional Biochemistry. 2002, 13(10): 625-633.
4. Hong Wang, Lisa A. Reaves, and Neilé K. Edens. Korean ginseng antilipolysis may be mediated in part by phosphodiesterase 4 in rat adipocytes in vitro. Obesity Research. 2003, 11(Suppl): A46.
viii 5. Hong Wang and Neilé K. Edens. Prostaglandin E2 antilipolysis may be mediated in part by phosphodiesterase 4 in rat adipocytes in vitro. Diabetes. 2004, 53(Suppl 2): A459.
6. Hong Wang and Neilé K. Edens. Expression of phosphodiesterase 4 in rat adipocytes. Obesity Research, 2004, 12(Suppl): A179.
FIELD OF STUDY
Major Field: The Ohio State University Nutrition Program
ix
TABLE OF CONTENTS
Abstract…………………………………………………………………………………....ii Dedication………………………………………………………………………………....v Acknowledgements……………………………………………………………………….vi Vita……………………………………………………………………………………...viii Table of contents………………………………………………………………………….x List of tables……………………………………………………………………………..xii List of figures……………………………………………………………………………xiv List of abbreviations…………………………………………………………………….xvi
Chapters: 1 Literature review……………………………………………………………………...1 1.1 Overview of type 2 diabetes……………………………………………………1 1.1.1 Insulin resistance…………………………………………………………..2 1.1.2 Impaired insulin secretion…………………………………………………3 1.2 Insulin signaling………………………………………………………………..4 1.2.1 Insulin signaling pathway………………………………………………….5 1.2.2 Abnormality of insulin signaling in type 2 diabetes………………………8 1.3 Adipose tissue…………………………………………………………………9 1.3.1 Adipose tissue distribution……………………………………………….10 1.3.2 Adipose tissue factors…………………………………………………….10 1.3.3 Regulation of lipolysis…………………………………………………...11 1.4 Effects of FFA on glucose metabolism………………………………………..19 1.4.1 Skeletal muscle…………………………………………………………...19 1.4.2 Liver……………………………………………………………………...21 1.4.3 Pancreatic β-cells………………………………………………………...23 1.4.4 Antilipolytic agents in treatment of type 2 diabetes……………………...24 1.5 Ginseng and type 2 diabetes…………………………………………………..25 1.5.1 Introduction to ginseng…………………………………………………...26 1.5.2 Ginsenosides……………………………………………………………...27 1.5.3 Hypoglycemic effect of ginseng………………………………………....29 1.5.4 Mechanisms for the hypoglycemic effect of ginseng……………………32 1.5.5 Summary…………………………………………………………………33 2 General methods…………………………………………………………………….41 2.1 Chemicals……………………………………………………………………..41 2.2 Animal………………………………………………………………………..42
x 2.3 Adipocyte isolation……………………………………………………………42 2.4 Lipolysis assay………………………………………………………………..43 2.5 Adipocyte RNA isolation……………………………………………………..44 2.6 Reverse transcription-polymerase chain reaction……………………………..44 2.7 Real-time quantitative reverse transcription polymerase chain reaction……...45 2.8 Western blot analysis………………………………………………………….46 2.8.1 Preparation of crude subcellular fractions………………………………..46 2.8.2 PKB phosphorylation…………………………………………………….46 2.8.3 PDE3B and PDE4B expression…………………………………………..47 2.9 Purification of [3H] cAMP…………………………………………………….48 2.9.1 Thin layer chromatography purification…………………………………48 2.9.2 Column purification……………………………………………………...49 2.10 PDE activity assay…………………………………………………………….49 2.11 Statistical analysis……………………………………………………………..50 3 Mechanisms for ginseng antilipolysis in rat adipocytes in vitro……………………53 3.1 Introduction……………………………………………………………………53 3.2 Effects of insulin and KGE on lipolysis………………………………………55 3.3 Effects of insulin and KGE on PKB phosphorylation and translocation……..56 3.4 Discussion……………………………………………………………………..58 4 The expression of phosphodiesterase 4 in rat adipocytes…………………………...67 4.1 Introduction……………………………………………………………………67 4.2 Determination of PDE4 gene expression in rat adipocytes…………………...70 4.3 Quantification of PDE4 gene expression in rat adipocytes…………………...71 4.4 PDE3B and PDE4B protein expression in rat adipocytes…………………….73 4.5 Discussion……………………………………………………………………..74 5 The role of phosphodiesterase 4 in antilipolysis mediated by PGE2 and adenosine..94 5.1 Introduction……………………………………………………………………94 5.2 Effects of insulin and PGE2 on lipolysis………………………………………95 5.3 Effects of insulin and PIA on lipolysis………………………………………..96 5.4 Effects of PDE inhibitors on basal lipolysis…………………………………97 5.5 Discussion……………………………………………………………………98 6 Activation of phosphodiesterase 4 in rat adipocytes in vitro………………………104 6.1 Introduction…………………………………………………………………..104 6.2 Determination of suitable concentrations of PDE3 and PDE4 inhibitors……105 6.3 Subcellular distribution of PDE4 activity in rat adipocytes…………………107 6.4 Effects of PGE2, PIA, and KGE on PDE activity in rat adipocytes……....108 6.5 Discussion……………………………………………………………………109 7 Conclusions………………………………………………………………………...117 Epilogue………………………………………………………………………………...120 Bibliography……………………………………………………………………………121
xi
LIST OF TABLES
Table Page
1.1 The regulation and function of PKC isoforms in insulin signaling………………...37
3.1 Comparison of ginsenoside profiles between AGE and KGE……………………...66
4.1 Comparison of PDE3 and PDE4……………………………………………………78
4.2 Primer pairs for RT-PCR to detect sequences encoding PDE3B and PDE4 (A, B, C, D)……………………………………………………………………………...... 79
4.3 Sequence of PDE3B PCR product aligned with the published sequence of PDE3B using NCBI two-sequence BLAST…………………………………………………81
4.4 Sequence of PDE4A PCR product aligned with the published sequence of PDE4A using NCBI two-sequence BLAST…………………………………………………82
4.5 Sequence of PDE4B PCR product aligned with the published sequence of PDE4B using NCBI two-sequence BLAST…………………………………………………83
4.6 Sequence of PDE4C PCR product aligned with the published sequence of PDE4C using NCBI two-sequence BLAST…………………………………………………84
4.7 Sequence of PDE4D PCR product aligned with the published sequence of PDE4D using NCBI two-sequence BLAST…………………………………………………85
4.8 Alignment of PDE3B and PDE4 (A, B, C, D) primers and probes for QPCR……..86
4.9 A region of high similarity among PDE3B and PDE4 (A, B, C, D) sequences……87
4.10 Properties of QPCR amplicons used for relative quantitation of PDE4 (A, B, C, D) gene expression……………………………………………………………………..89
xii 6.1 Effects of different concentrations of the PDE3 and PDE4 inhibitors on PDE3 and PDE4 activity in rat adipocytes……………………………………………………113
6.2 Effects of various agents on PDE3 and PDE4 activity in rat adipocytes………….116
xiii
LIST OF FIGURES
Figure Page
1.1 Schematic representation of insulin signaling……………………………………...35
1.2 Schematic representation of regulation of lipolysis………………………………...36
1.3 Biosynthesis of prostagladins………………………………………………………38
1.4 The chemical structures of ginsenosides……………………………………………39
2.1 The principle for the Taqman method of QPCR……………………………………52
3.1 Comparison of AGE and KGE antilipolysis in rat adipocytes………………….62
3.2 Effects of insulin and KGE on lipolysis in rat adipocytes………………………….63
3.3 Effects of insulin and a range of concentrations of KGE on PKB phosphorylation in cell lysates of rat adipocytes………………………………………………………64
3.4 Effects of insulin and KGE on PKB phosphorylation in cell lysates, cytosolic fraction, and particulate fraction of rat adipocytes…………………………………65
4.1 Expression of PDE3B and PDE4 (A, B, C, D) in rat adipocytes…………………...80
4.2 The standard curves for the amplification of PDE3B and PDE4 (A, B, C, D)……..88
4.3 Relative expression of PDE3B and PDE4 (A, B, C, D) normalized to 28s RNA in rat adipocytes…………………………………………………………………………..90
4.4 Relative expression of PDE3B and PDE4 (A, B, C, D) normalized to 28s RNA in rat heart tissue………………………………………………………………………….91
4.5 Expression of PDE3B in rat adipocytes…………………………………………….92
4.6 Expression of PDE4B3 in rat adipocytes…………………………………………...93
xiv
5.1 Effects of insulin and PGE2 on lipolysis in rat adipocytes………………………101
5.2 Effects of insulin and PIA on lipolysis in rat adipocytes………………………102
5.3 Effects of the PDE3 and PDE4 inhibitors on basal lipolysis in rat adipocytes…103
6.1 Subcellular distribution of total PDE, PDE3B, and PDE4 activity in rat adipocytes assayed at 0.1 µM cAMP…………………………………………………………114
6.2 Effects of various agents on the total PDE activity in rat adipocytes……………..115
xv
LIST OF ABBREVIATIONS
PDE Phosphodiesterase PKB Protein kinase B PI3-K Phosphatidylinositol 3–kinase KGE Korean ginseng extract AGE American ginseng extract FFA Free fatty acids cAMP cyclic adenosine monophosphate
PGE2 Prostaglandin E2 PIA N6-(2-phenylisopropyl)adenosine ISO Isoproterenol RT-PCR Reverse transcription-polymerase chain reaction QPCR Real-time quantitative reverse transcription polymerase chain reaction
xvi
CHAPTER 1
LITERATURE REVIEW
1.1 Overview of type 2 diabetes
Type 2 diabetes mellitus (Type 2 diabetes) is a metabolic syndrome characterized by hyperglycemia. Insulin resistance and impaired insulin secretion are two pathological characteristics in type 2 diabetes (Bergman et al., 2002; DeFronzo et al., 1992; Gerich,
2002). In the United States, it is estimated that over 16 million people are affected by diabetes mellitus. Type 2 diabetes accounts for 90-95% of all diabetes (Skyler and Oddo,
2002). The causes of type 2 diabetes include genetic and environmental factors (Hamman,
1992). Until now, only a few specific genetic abnormalities in type 2 diabetes have been identified, such as maturity-onset diabetes of young (MODY) (Velho and Froguel, 1998).
The range of environmental factors is broad, including age, gender, ethnicity, physical activity, diet, smoking, obesity, and fat distribution (Gerich, 1998). Among these environmental factors, obesity has received more attention in recent years. The majority of type 2 diabetic patients are overweight or obese. According to the data provided by
Behavioural Risk Factor Surveillance System (BRFSS), there was a significant
1 correlation (r = 0.64, p<0.001) between the incidence of diabetes and the incidence of obesity among approximately 150,000 people between 1990 and 1998 in the USA
(Mokdad et al., 2000).
1.1.1 Insulin resistance
Insulin resistance is defined as the impaired ability of insulin to appropriately stimulate glucose disposal or suppress endogenous glucose production (McGarry, 2002;
Summers et al., 2000). Insulin resistance is associated with obesity, hypertension, dyslipidemia, and other conditions (Cefalu, 2001). Liver, skeletal muscle, and adipose tissue are the major sites for insulin resistance (DeFronzo et al., 1992). Insulin resistance is characterized by decreased glucose uptake and utilization by skeletal muscle, increased glucose production in liver, and increased lipolysis in adipose tissue (Bergman and Ader,
2000). The most widely used “gold standard” for assessing insulin resistance is the euglycemic hyperinsulinemic clamp. With this method, insulin is infused intravenously to achieve a certain plasma insulin level. Simultaneously, glucose is infused to maintain normal glycemia. The amount of glucose infused is a measurement of insulin sensitivity
(Cefalu, 2001).
One prevalent view is that insulin resistance is the primary factor in the pathogenesis of type 2 diabetes. Many long-term studies in different ethnic populations have shown that hyperinsulinemia precedes the onset of type 2 diabetes for many years (Martin et al.,
1992; Saad et al., 1989; Sicree et al., 1987). DeFronzo et al. proposed that insulin resistance exists first, thereby causing β-cells to secrete more insulin to compensate for insulin resistance. Hyperinsulinemia and normoglycemia are two features of this stage.
2 Sustained high demand for insulin eventually leads to β-cell dysfunction. When β-cells fail to secrete sufficient insulin to maintain normoglycemia, clinically evident type 2 diabetes occurs (DeFronzo et al., 1992).
1.1.2 Impaired insulin secretion
Impaired insulin secretion is another characteristic of type 2 diabetes. In the normal physiological state, insulin is secreted in a biphasic pattern in response to intravenous glucose challenge. An early phase of insulin secretion occurs within the first 10 minutes, and then a late phase of insulin secretion persists until the hyperglycemic stimulus disappears. The amount of insulin released from pancreatic β-cells during the early phase and the late phase is approximately 2 to 3% and 20% of the total insulin content, respectively (Del Prato, 2003). This biphasic pattern of insulin secretion is not obvious in response to an oral glucose load because plasma glucose concentration rises gradually
(DeFronzo, 2004).
The fasting plasma insulin concentration is normal or increased in type 2 diabetic patients. However, the early phase of insulin secretion is significantly reduced during oral glucose tolerance test (0-30 min) and during intravenous glucose tolerance test (0-10 min) in type 2 diabetes (DeFronzo, 2004). The early phase of insulin secretion plays a critical role in the maintenance of postprandial glucose homeostasis mainly by suppressing endogenous glucose production (EGP). Thus, loss of early insulin secretion leads to postprandial hyperglycemia and hyperinsulinemia (Del Prato, 2003).
In contrast with the above view about the primacy of insulin resistance, others have proposed that impaired insulin secretion is the primary defect in the pathogenesis of type
3 2 diabetes. Supportive evidence for this hypothesis is the significant reduction in the early
phase of insulin secretion in severely insulin-resistant nondiabetic offspring of type 2 diabetic parents (Perseghin et al., 1997). Some researchers proposed that the dysfunctional β-cells already exist in the preclinical stage, but β-cells can secrete enough
insulin to maintain normoglycemia. Progression to type 2 diabetes results from further
deterioration of insulin secretion and increased insulin resistance (Kahn, 2003; Ostenson,
2001).
Taken together, insulin resistance is an early event in type 2 diabetes and impaired
insulin secretion is a prerequisite for progression to type 2 diabetes. Although the relative
primacy of insulin resistance and impaired insulin secretion is still debatable, it is
generally recognized that both these factors play important roles in the pathogenesis of
type 2 diabetes.
1.2 Insulin signaling
Defects in insulin signaling play a role in the insulin resistance of type 2 diabetes at
the cellular level and whole-body level (DeFronzo, 2004). It is well known that insulin is
a potent anabolic hormone essential for maintaining whole-body glucose homeostasis and
regulating many other physiological functions (Rhodes and White, 2002; Zick, 2001).
Insulin increases glucose uptake and utilization in skeletal muscle and adipose tissue, and
decreases endogenous glucose production and promotes glycogen synthesis in liver.
Insulin also increases lipid synthesis in liver and adipose tissue, and inhibits fatty acid
release from triglycerides in adipose tissue (Saltiel and Kahn, 2001). In addition, insulin
promotes protein synthesis and regulates the expression of many genes (Rhodes and
4 White, 2002). The diverse actions of insulin are coordinated through a complex signaling
system (Figure 1.1). Defects in insulin signaling impair insulin action in insulin-sensitive
tissues, thus adversely affecting the above physiological functions regulated by insulin.
1.2.1 Insulin signaling pathway
Insulin binding to the α-subunit of its receptor activates the tyrosine kinase activity
of the β-subunit, leading to autophosphorylation of tyrosine residues in the receptor
β-subunit. This autophosphorylation increases tyrosine kinase activity of the insulin
receptor. Subsequently, the tyrosine kinase activity of the insulin receptor phosphorylates
a series of tyrosine residues in protein substrates, including the insulin receptor substrate
proteins (IRS1/2/3/4), Src homology 2 containing cytoplasmic adapter protein (Shc),
signal regulated protein (SIRP), growth factor receptor-binding protein associated
binder-1 (Gab-1), Cbl, and adapter protein with a PH and SH2 domain (APS) (Pessin and
Saltiel, 2000; Rhodes and White, 2002; Saltiel and Kahn, 2001).
Tyrosine phosphorylation of the IRS proteins provides the binding sites for the
downstream molecules containing Src homology 2 (SH2) domains, including growth
factor receptor binding protein 2 (Grb2), SH2-containing protein tyrosine phosphatase-2
(SHP-2), and phosphatidylinositol 3–kinase (PI3-K). These downstream signaling
molecules are activated by binding to IRS (Pessin and Saltiel, 2000; Rhodes and White,
2002; Saltiel and Kahn, 2001).
PI3-K is a critical signaling molecule that mediates diverse actions of insulin in a variety of cell types. PI3-K is composed of two subunits, a p110 catalytic subunit and a p85 regulatory subunit containing two SH2 domains (Myers et al., 1992). The interaction 5 of IRS proteins with the p85 subunit of PI3-K activates the p110 subunit of PI3-K,
leading to the conversion of phosphatidylinositol 4,5-bisphosphate (PIP2) to
phosphatidylinositol 3,4,5-trisphosphate (PIP3) within plasma membrane. Subsequently,
PIP3 activates 3-phosphoinositide-dependent protein kinase-1 (PDK-1), which mediates
the phosphorylation of the downstream signaling molecules, including protein kinase B
(PKB, also known as Akt), atypical protein kinase C isoforms (PKCζ/λ), and p70
ribosomal protein S6 kinase (p70S6K) (Farese, 2001).
In skeletal muscle and adipose tissue, PI3-K mediates insulin-stimulated glucose
uptake and GLUT4 translocation by activation of the downstream targets, PKB and
atypical PKCζ/λ (Czech and Corvera, 1999) (Pessin and Saltiel, 2000). PKC isoforms are
divided into 3 groups: classical PKC (cPKC), novel PKC (nPKC), and atypical PKC
(aPKC). In contrast with aPKC isoforms, cPKC and nPKC isoforms can be activated by
diacylglycerol (DAG) and downregulate insulin signaling (Jiang and Zhang, 2002)
(Table 1.1). In addition, the insulin receptor also phosphorylates the Cbl/CAP complex.
Once phosphorylated, the Cbl/CAP complex translocates to lipid rafts in plasma
membrane and recruits the adapter proteins CrK and C3G, leading to translocation and
activation of the G protein TC10 by exchanging GTP for GDP. Activation of TC10
promotes GLUT4 translocation to plasma membrane (Saltiel and Kahn, 2001).
In liver and muscle, insulin stimulates glycogen synthesis through
dephosphorylation and activation of glycogen synthase. PKB, the downstream target of
PI3-K, phosphorylates and inactivates glycogen synthase kinase, limiting
phosphorylation of glycogen synthase and thus increasing its activity (Cross et al., 1995).
In liver, insulin also regulates the expression of genes responsible for glycolysis and 6 gluconeogenesis. Insulin up-regulates the expression of genes encoding glycolytic
enzymes (e.g. glucokinase and pyruvate kinase), and down-regulates the expression of genes encoding gluconeogenic enzymes (e.g. phosphoenolpyruvate carboxykinase and
glucose 6-phosphatase) (Saltiel and Kahn, 2001). IRS-2 plays a key role in insulin signal
transduction in liver. In primary hepatocytes isolated from IRS-2(-/-) mice, insulin did not
inhibit the expression of gluconeogenic genes (Valverde et al., 2003).
In adipose tissue, PI3-K also plays a key role in insulin-mediated antilipolysis
through phosphorylation and activation of phosphodiesterase 3B (PDE3B) (Degerman et
al., 1998). Wortmannin, a selective PI3-K inhibitor, inhibits phosphorylation and activation of PDE3B and the antilipolytic effect of insulin (Rahn et al., 1994). The downstream kinase activated by PI3-K responsible for phosphorylating and activating
PDE3B is possibly PKB rather than mitogen-activated protein kinase or p70S6K
(Wijkander et al., 1998).
Insulin increases lipogenesis in adipocytes by increasing the uptake of glucose and activating lipogenic (e.g. acetyl-CoA carboxylase, fatty acid synthase and glycerol-P acyltransferase) and glycolytic enzymes (e.g. glucokinase and pyruvate kinase). Insulin also has a long-term effect on lipogenesis by up-regulating the expression of lipogenic genes, particularly sterol regulatory element binding protein-1 (SREBP-1) in liver and peroxisome proliferator-activated receptor γ (PPARγ) in adipose tissue (Kersten, 2001).
Insulin stimulates protein synthesis through activation of the mammalian target of rapamycin (mTOR) which is a downstream signaling molecule of PI3-K. After being activated by PI3-K, mTOR phosphorylates and activates p70S6K to increase translation of mRNA (Rhodes and White, 2002; Saltiel and Kahn, 2001).
7 Insulin regulates gene expression by stimulating the mitogen-activated protein kinase (MAPK) cascades. Phosphorylation of tyrosine in IRS recruits the adapter protein
Grb2 which interacts with the Son of the sevenless (SOS) exchange protein to activate
Ras. The activation of Ras elicits a series of activations including Raf, MAPK/ERK kinase (MEK) and extracellular signal-regulated kinase (ERK). Activated ERK
translocates into the nucleus and regulates gene expression (Rhodes and White, 2002;
Saltiel and Kahn, 2001).
1.2.2 Abnormality of insulin signaling in type 2 diabetes
The studies of various tissue-specific knockout models in animals and the ob/ob
mouse model (genetic deficiency of leptin) have shown that defects in insulin signaling
are correlated with insulin resistance. The muscle-specific insulin receptor-knockout
mouse was characterized by increased fat mass, serum triglycerides and FFA, but normal
blood glucose, serum insulin, and glucose tolerance (Bruning et al., 1998). This study
suggested that insulin resistance in muscle is associated with abnormal lipid metabolism
in type 2 diabetes, and that other organs and tissues contribute more to insulin-regulated
glucose disposal. The liver-specific insulin receptor-knockout mouse showed high
plasma insulin, glucose intolerance, and inability of insulin to inhibit hepatic glucose
production, suggesting the important role of insulin signaling in liver in maintaining
glucose homeostasis (Michael et al., 2000). PKBβ-knockout mice displayed decreased
insulin-stimulated glucose uptake in muscle and failure of insulin to suppress hepatic
glucose production in liver, coupled with increased pancreatic islet mass. These data
indicate that PKBβ is required for the maintenance of glucose homeostasis in mice (Cho
8 et al., 2001). Also, there was a remarkable decrease in insulin-stimulated IRS-1
phosphorylation and PI3-K activity in liver and muscle of the ob/ob mice, which may
contribute to insulin resistance (Folli et al., 1993).
A number of defects in insulin signaling have been demonstrated in muscle biopsies
of human subjects. It was reported that phosphorylation of the insulin receptor and IRS-1, and insulin-stimulated association of the p85 regulatory subunit of PI3-K with IRS-1 was significantly decreased in muscle biopsies of obese nondiabetic subjects and absent in type 2 diabetic subjects (Cusi et al., 2000). Another study also demonstrated that IRS-1 phosphorylation, PI3-K activity, and glucose transport activity were significantly reduced in skeletal muscle biopsies of type 2 diabetic subjects compared to lean healthy subjects
(Krook et al., 2000). A study in isolated human adipocytes from the lower abdominal region demonstrated that insulin-stimulated tyrosine kinase activity was reduced approximately 50% in type 2 diabetic patients compared to lean and obese nondiabetic subjects (Freidenberg et al., 1987). Another study in isolated human hepatocytes from intraoperative liver biopsies also reported that insulin-stimulated tyrosine kinase activity was decreased in diabetic subjects (Caro et al., 1986).
In summary, insulin signaling is necessary for normal metabolism of glucose, lipid, and protein, as well as growth and development. Abnormal insulin signaling in target organs and tissues is responsible for the insulin resistance in type 2 diabetes.
1.3 Adipose tissue
Traditionally, adipose tissue is regarded as a major site for energy storage. Fat is stored as triglyceride (TG) in adipose tissue, and is mobilized from adipose tissue as FFA
9 when fuel is needed. Adipose now is also recognized as an endocrine organ (Ahima and
Flier, 2000). The factors secreted from adipose tissue have diverse effects on insulin
signaling.
1.3.1 Adipose tissue distribution
Adipose tissue includes visceral adipose tissue and subcutaneous adipose tissue. It is
thought that visceral adipose tissue contributes more to insulin resistance than
subcutaneous adipose tissue (Bergman et al., 2001). First, compared to subcutaneous
adipocytes, visceral adipocytes have a high rate of basal lipolysis and are more
responsive to lipolytic hormones (e.g. catecholamines), but less responsive to
antilipolytic hormones (e.g. insulin) (Raz et al., 2005). Second, visceral adipose tissue
directly delivers FFA to liver through the portal vein. Enlarged visceral adipose tissue
results in high FFA exposure in liver, leading to increased endogenous glucose production
(Bergman et al., 2001).
1.3.2 Adipose tissue factors
Adipose tissue releases many factors with diverse functions into blood stream. Some of these are known to affect whole-body glucose metabolism. These factors include FFA, leptin, tumor necrosis factor-α (TNF-α), interleukin-6 (IL-6), adiponectin, and resistin.
FFA will be discussed in detail in Section 1.4. Leptin acts on its receptors in the central
nervous system to decrease food intake and increase energy expenditure. Expression of
TNF-α gene is increased in obesity. TNF-α increases insulin resistance by inducing
lipolysis in an autocrine and paracrine manner. IL-6 is a circulating cytokine. Plasma 10 IL-6 concentration is positively correlated with insulin resistance. Expression of
adiponectin mRNA is reduced in obesity. Upregulation of adiponectin may ameliorate
insulin resistance. Resistin is a newly discovered polypeptide that is related to the
development of insulin resistance (Pittas et al., 2004).
1.3.3 Regulation of lipolysis
The hydrolysis of stored TG is called lipolysis. Lipolysis is catalyzed by enzymes
and yields FFA and glycerol. It was once thought that hormone-sensitive lipase (HSL)
was the only enzyme to hydrolyze TG. Recently, three independent laboratories reported
that a second enzyme, adipose triglyceride lipase (ATGL), also specifically hydrolyzes
TG (Jenkins et al., 2004; Villena et al., 2004; Zimmermann et al., 2004). The mechanism
by which HSL regulates lipolysis is relatively clear. Lipolysis is mediated by increased intracellular concentration of cyclic adenosine monophosphate (cAMP). cAMP is synthesized by adenylate cyclase and degraded by phosphodiesterases. The increase in
cAMP concentration leads to activation of cAMP-dependent protein kinase A (PKA)
(Holm et al., 2000). When the cAMP concentration is low, PKA exists as an inactive
heterotetrameric complex containing two regulatory subunits and two catalytic subunits.
When the cAMP concentration is increased, it binds to the regulatory subunits, and the
catalytic units are released in an active form (Michel and Scott, 2002). Ultimately, the
PKA target proteins HSL and perilipin A are phosphorylated and the rate of lipolysis is
dramatically increased (Holm et al., 2000).
The regulation of lipolysis in adipocytes is complex and affected by many factors
(Figure 1.2). Catecholamines and insulin are classical lipolytic and antilipolytic
11 hormones, repectively. Adenosine and prostaglandin E2 are secreted by adipocytes and
exert their antilipolytic action in an autocrine and paracrine manner. The phosphorylation state of hormone-sensitive lipase and perilipin A determine the rate of lipolysis.
1.3.3.1 Hormone-sensitive lipase
Hormone-sensitive lipase (HSL) catalyzes the hydrolysis of triglyceride to
diglyceride and in turn to monoglyceride in a stepwise manner. The first step is
rate-limiting in breakdown of triglyceride, so HSL is the rate-limiting enzyme (Langin et al., 1996). HSL can be phosphorylated at Ser-563, Ser-565, Ser-659, and Ser-660. Ser-563 and Ser-565 are two sites first identified to be phosphorylated in HSL. Initially, Ser-563 was named as the “regulatory site”, because it is phosphorylated by PKA in response to
increase in cAMP; Ser-565 was named the “basal site”, because it is found to be
phosphorylated not by PKA but by other kinases, including AMP-activated protein kinase
(AMPK), Ca2+/calmodulin-dependent kinase, and glycogen synthase kinase.
Phosphorylation of Ser-563 and phophorylation of Ser-565 seem mutually exclusive.
Therefore, phophorylation of Ser-565 by AMPK may inhibit catecholamine-induced lipolysis (Londos et al., 1999).
In recent years, two other PKA-phosphorylation sites, Ser-659 and Ser-660, were reported to be responsible for HSL activation in vitro (Anthonsen et al., 1998). Recent
mutational analysis demonstrated that Ser-659 and Ser-660 are involved in translocation
and activation of HSL (Su et al., 2003). Evidence from subcellular fractionation in rat
adipocytes (Clifford et al., 2000) and immunofluorescene microscopy in 3T3-L1
adipocytes (Brasaemle et al., 2000a) showed that HSL was scattered throughout the
12 cytosol in the basal state and HSL translocated from the cytosol to the surface of lipid droplets upon catecholamine stimulation, thus initiating lipolysis.
Alterations in the HSL gene may contribute to the genetic predisposition to type 2 diabetes. It was reported that a significant difference in allele frequency distribution in the human HSL gene using a polymorphic marker between type 2 diabetic patients and control subjects, particularly type 2 diabetic patients with abdominal obesity
(Klannemark et al., 1998). HSL knock-out study showed the HSL-deficient mice had reduced circulating FFA concentration and intramuscular TG content, and were prevented from diet-induced insulin resistance in skeletal muscle compared to the wild-type mice after high fat feeding (Park et al., 2005).
1.3.3.2 Perilipin
Perilipins are a family of related proteins located on the surface of the lipid droplets in adipocytes. Perilipin A is the most abundant isoform in adipocytes (Londos et al.,
1999). Perilipin A covers the surface of the lipid droplet and inhibits HSL binding to the lipid droplet in the basal state. Therefore, it is considered a tonic inhibitor of basal lipolysis (Brasaemle et al., 2000b). Phosphorylation of perilipin A by PKA at Ser-81,
Ser-222, and Ser-276 makes perilipin A disassociate from the lipid droplet and simultaneously allows phosphorylated HSL to access the lipid droplet, leading to initiation of lipolysis (Souza et al., 2002). More recent results showed that perilipin A is also required for the translocation of HSL during stimulated lipolysis (Sztalryd et al.,
2003).
13 The expression of perilipin A in obesity is not consistent. Perilipin A content
decreased 50% in subcutaneous adipocytes isolated from obese women compared to
nonobese women, and basal and norepinephrine-induced lipolysis increased 2 to 4-fold.
There was an inverse correlation between perilipin A expression and the concentrations
of circulating glycerol and FFA (Mottagui-Tabar et al., 2003). In contrast with this study,
perilipin A expression in abdominal subcutaneous adipose tissue was significantly correlated with obesity (r = 0.55; P < 0.01, perilipin A mRNA vs. percent body fat).
However, no significant correlation was found between perilipin A expression and plasma FFA, and between perilipin A expression and insulin resistance (Kern et al.,
2004). The reason for this discrepancy is not clear.
1.3.3.3 Catecholamines
The catecholamines include epinephrine and norepinephrine. The adrenal medulla releases mainly epinephrine into the circulation, and sympathetic nerves releases mainly norepinephrine to adipose tissues in response to stress. Therefore, lipolysis in adipose tissue is regulated by catecholamines systemically and locally (Rayner, 2001).
Catecholamines regulate lipolysis via four adrenergic receptors: β1, β2, β3, and α2
(Lafontan et al., 1995). All three β-adrenergic receptors (β-ARs) couple to the stimulatory
G-protein known as Gs and activate adenylate cyclase, increasing cAMP concentration
and stimulating lipolysis. However, β3-ARs can also interact with inhibitory G-protein known as Gi in adipocytes (Granneman, 1995). β1-ARs can be activated by
catecholamines at low concentrations; β3-ARs require higher concentration for activation
(Galitzky et al., 1993; Lafontan et al., 1995). In contrast with β-ARs, α2-adrenergic 14 receptors (α2-ARs) couple Gi and inhibit adenylate cyclase, decreasing cAMP
concentration and suppressing lipolysis (Lafontan and Berlan, 1995). The distribution of
adrenergic receptor subtypes determines the relative efficacy of catecholamines for
lipolysis in adipocytes (Lafontan and Berlan, 1993).
In the condition of hyperinsulinemia caused by hyperinsulinemic euglycemic
clamp in healthy male subjects, epinephrine-induced lipolysis was significantly
inhibited during the clamp. When epinephrine was infused together with
phentolamine (a nonselective α-AR antagonist), the inhibition of epinephrine-induced
lipolysis during hyperinsulinemic euglycemic clamp was reduced significantly (Stich et al., 2003). This study suggests that activation of α2-ARs by epinephrine is involved in inhibition of epinephrine-induced lipolysis by hyperinsulinemia in subcutaneous adipose tissue.
1.3.3.4 Insulin
Insulin inhibits lipolysis by phosphorylation and activation of phosphodiesterase 3B
(PDE3B) (Degerman et al., 1990; Smith et al., 1991). PDE3B is the major isoform of
PDE in adipose tissue. PDE3B can degrade cAMP to AMP, thus decreasing intracellular cAMP concentration. In isolated rat adipocytes, the specific PDE3B inhibitors cilostamide and OPC 3911 can reverse the antilipolytic effect of insulin (Elks and
Manganiello, 1984; Eriksson et al., 1995; Schmitz-Peiffer et al., 1992). Insulin stimulation phosphorylates PDE3B at Ser-302 in rat adipocytes (Rahn et al., 1996) and at
Ser-273 in a cultured adipocyte line derived from mouse possibly as a result of species specificity (Kitamura et al., 1999). The activation of PDE3B enhances degradation of 15 intracellular cAMP, leading to decrease in PKA activity, concomitant reduction of HSL
activity, and a decrease in the rate of lipolysis (Degerman et al., 1998).
A previous study in human subcutaneous adipose tissue from diabetic patients reported a significant decrease in PDE3B activity that was normalized after treatment with insulin or oral antidiabetic agents (Engfeldt et al., 1982). TNF-α-induced lipolysis was shown to be mediated by down-regulating PDE3B expression in murine 3T3-L1 adipocytes (Rahn Landstrom et al., 2000) and differentiated human adipocytes (Zhang et al., 2002).
1.3.3.5 Adenosine
Adenosine is released from adipocytes and suppresses lipolysis by inhibiting adenylate cyclase (Fain and Wieser, 1975; Vassaux et al., 1992). Adenosine has three receptor subtypes: A1, A2, and A3. A1 and A3 inhibit adenylate cyclase; and A2 (A2a and
A2b) activates adenylate cyclase (Yaar et al., 2005). Adenosine inhibits lipolysis in
adipocytes by binding to A1 adenosine receptors (A1-ARs) (Londos et al., 1978).
Adenosine is continuously released from adipocytes to the medium during incubation of
adipocytes (Fain and Wieser, 1975). Addition of adenosine deaminase into incubation
medium hydrolyzes endogenous adenosine, leading to cAMP accumulation and increased
lipolysis (Honnor et al., 1985). A1-ARs are highly expressed in rat adipocytes and are very sensitive to the agonist adenosine, and can be activated by endogenous adenosine at nanomolar concentration (Liang et al., 2002).
Adenosine has been demonstrated to enhance insulin action through increasing the insulin-induced production of PIP3 by PI3-K and activation of PKB in rat adipocytes
16 (Takasuga et al., 1999). In transgenic mice overexpressing A1-ARs in adipose tissue, lower plasma FFA level was observed compared to the control mice, suggesting an important role for adipocyte A1-AR in the regulation of lipolysis. It is more interesting to
note that a high fat diet did not induce insulin resistance on the transgenic mice measured by fasting serum glucose and insulin levels, as well as glucose and insulin tolerance tests, although both the transgenic mice and the control mice developed obesity. This study suggests that overexpression of A1-ARs in murine adipocytes prevents insulin resistance
associated with obesity (Dong et al., 2001).
1.3.3.6 Prostaglandin E2
Prostaglandins (PGs) are major products derived from arachidonic acid (AA).
Various stimuli increase phospholipase A2 (PL A2)-mediated hydrolysis of membrane
phospholipids to form AA. Sequentially, AA is converted to PGH2 including an unstable
intermediate PGG2 by cyclooxygenase (COX). PGH2 is converted to a series of PGs by
respective PG synthases depending on the cell-specific expression of PG synthases. Until
now, five PGs have been identified, including PGE2, PGD2, PGF2α, PGI2, and
thromboxane A2 (TXA2) (Figure 1.3). These PGs exert their multiple physiological
functions in an autocrine and paracrine manner. Some of the major functions of PGs
include relaxation and contraction of smooth muscles, secretion and motility of
gastrointestinal tract, and the transport of ions and water in kidney (Narumiya et al.,
1999).
PGE2 is one of the PGs released from adipocytes (Richelsen and Pedersen, 1987;
Shaw and Ramwell, 1968). PGE2 is the only prostaglandin that has a clear antilipolytic
17 effect (Vassaux et al., 1992). PGE2 has 4 receptor subtypes, known as EP1, EP2, EP3, and
2+ EP4. Among them, the EP1 receptor mediates PGE2-induced increases in free Ca concentration via an unidentified G protein. The EP2 and EP4 receptors increase cAMP
concentration by activating adenylate cyclase via Gs activation. The EP3 receptor decreases cAMP concentration by inhibiting adenylate cyclase via Gi activation
(Narumiya et al., 1999). EP1, EP3, and EP4 receptor mRNA have been identified in
mature adipocytes. The expression of EP1 and EP4 mRNA is predominant in
preadipocytes, and EP3 mRNA is exclusively expressed in mature adipocytes (Borglum et
al., 1999). The role of EP1 and EP4 in mature adipocytes is not clear, and EP3 is involved
in inhibition of lipolysis by activating Gi protein, inhibiting adenylate cyclase, and
decreasing cAMP (Borglum et al., 1999; Strong et al., 1992).
The antilipolytic effect of PGE2 was significantly reduced in obese subjects compared to nonobese subjects due to the reduction of PGE2 binding relative to adipocyte
surface area (Richelsen, 1988). A previous study compared the effects of lipolytic and
antilipolytic agents on lipolysis in adipocytes isolated from various fat depots in females.
The results demonstrated that epinephrine had minimal lipolytic effect in adipocytes from
the subcutaneous regions, but significantly increased lipolytic effect in omental adipocytes. This difference was mainly due to the predominant expression of α2-AR in subcutaneous adipocytes. In contrast, the antilipolytic effects of insulin and clonidine
(α2-AR agonist) were significantly less in omental adipocytes compared to subcutaneous adipocytes. However, PGE2 had similar antilipolytic effect in adipocytes from the various
depots. These data suggest that omental adipose tissue is specifically resistant to insulin,
but not PGE2 (Richelsen et al., 1991). 18 1.4 Effects of FFA on glucose metabolism
FFA are released from adipocytes during lipolysis. Elevated plasma FFA
concentration is common in obese people (Golay et al., 1986; Jansson et al., 1992) and
people with type 2 diabetes (Groop et al., 1989; Reaven et al., 1988). The possible
reasons for high FFA levels include enlarged adipose tissue mass and increased rates of
basal and stimulated lipolysis (Arner, 2002; Bergman et al., 2001; Raz et al., 2005). High
circulating FFA concentration impairs insulin action and glucose homeostasis. Increasing plasma FFA levels acutely by intravenous lipid infusion delayed disappearance of infused
6,6 D2-glucose disappearance for 2-4 h in healthy subjects and this defect was associated
with a decrease in muscle glycogen synthesis (Boden et al., 1991). Also, an acute
increase in plasma FFA levels generated a 40-50% decrease in whole body
insulin-stimulated glucose uptake determined with [3H] glucose in diabetic subjects
(Boden and Chen, 1995). Many studies have shown that chronically elevated plasma FFA
and concomitant lipid accumulation in insulin-responsive tissues cause insulin resistance
in muscle and liver and impair insulin secretion from pancreas (Bays et al., 2004; Boden,
1997; Carpentier et al., 2000; Kashyap et al., 2003; Kelley and Mandarino, 2000;
McGarry, 2002).
1.4.1 Skeletal muscle
Skeletal muscle is the major site for glucose uptake and utilization in response to
insulin stimulation. Also, it is the major site for β-oxidation of FFA. It is thought that
increased availability of FFA can interfere with glucose utilization in skeletal muscle
(Bergman and Ader, 2000). The development of the new technology, such as nuclear
19 magnetic resonance spectroscopy (NMR), allows researchers to study the lipid
environment of skeletal muscle in vivo (Shulman, 1999). For example, application of this technology has shown that increased intramyocellular triglyceride content is associated with insulin resistance in the offspring of subjects with type 2 diabetes (Perseghin et al.,
1999).
In 1960s, Randle proposed his glucose-fatty acid hypothesis to explain how FFA interfered with glucose utilization (Randle et al., 1963). Based on this hypothesis,
elevated plasma FFA concentration results in increased uptake of FFA. Excessive FFA
undergo β-oxidation in mitochondria, leading to an accumulation of acetyl-CoA.
Increased acetyl-CoA (or increased acetyl-CoA/CoA ratio) inhibits pyruvate
dehydrogenase. Also, increased formation of acetyl-CoA from FFA increases the muscle citrate which inhibits phosphofructokinase. Taken together, increased acetyl-CoA and
citrate levels therefore reduce the rate of glycolysis. The resultant accumulation of glucose-6-phosphate in the muscle inhibits hexokinase activity and thus reduces muscle
glucose uptake. However, contrary to Randle’s hypothesis, the reduced intracellular
glucose-6-phosphate concentration was reported after lipid infusion using magnetic resonance spectroscopy in healthy human subjects (Roden et al., 1996). Therefore,
Randle’s hypothesis appears to be insufficient to explain the effects of FFA in skeletal muscle.
A recent review (Ruderman et al., 1999) suggests that increased long-chain fatty acyl-CoA (LCFA-CoA) may interfere with glucose metabolism in skeletal muscle.
Increased availability of FFA causes an accumulation of acetyl-CoA by β-oxidation, and acetyl-CoA is in turn converted to malonyl-CoA by acetyl-CoA carboxylase. This 20 resultant increase in malonyl-CoA concentration inhibits carnitine palmitoyl transferase
(CPT-1). Because CPT-1 is responsible for transporting FFA into mitochondria for oxidation, inhibition of CPT-1 results in accumulation of LCFA-CoA in cytosol
(Jeukendrup, 2002). There are two possible explanations for the interference of
LCFA-CoA on glucose metabolism. First, LCFA-CoA inhibits glycogen synthase by dissociating the tetrameric enzyme into monomers and binding to the monomers
(Wititsuwannakul and Kim, 1977). Second, LCFA-CoA and its derivatives, diacylglycerol (DAG) and ceramide, activate novel PKCθ which phosphorylates IRS on serine/threonine residues of IRS. PKCθ-mediated phosphorylation of IRS subsequently decreases PI3-K activity (Shulman, 1999) (Table 1.1). It was showed that PKCθ was translocated to the plasma membrane of rat after lipid infusion and reduced the PI3-K activity associated with IRS-1 by more than 50% (Griffin et al., 1999).
1.4.2 Liver
Liver is a critical organ in the regulation of blood glucose level. Approximately 85% and 15% of endogenous glucose production (EGP) is from liver and kidney, respectively
(DeFronzo, 2004). In type 2 diabetes, high fasting blood glucose is mostly attributable to increased EGP (DeFronzo et al., 1992). EGP derives from gluconeogenesis and glycogenolysis (Cherrington, 1999). Gluconeogensis is synthesis of glucose from non-hexose precursors, such as pyruvate, amino acids, and glycerol. Glycogenolysis is breakdown of glycogen to glucose. The stimulation of gluconeogenesis may be a primary event responsible for hyperglycemia in type 2 diabetes. It was reported that a 60% increase in the rate of gluconeogenesis may account for the increased rates of glucose 21 production in type 2 diabetes by using 13C NMR spectroscopy to measure the rates of net hepatic gluconeogenesis and glycogenolysis. (Magnusson et al., 1992).
FFA are an important signal to regulate EGP, and thereby increased flux of FFA into liver results in overproduction of glucose (Bergman and Ader, 2000). The mechanisms by which FFA increase EGP are still unclear. One possible mechanism is that FFA stimulate
EGP through upregulating glucose-6-phosphatase expression in vivo (Massillon et al.,
1997). Another mechanism is that increased FFA concentration impairs insulin signaling in liver. A recent study in rats showed that the metabolites of FFA, VIZ., LCFA-CoA and
DAG, induced hepatic insulin resistance by the activation and translocation of novel
PKC-δ to plasma membrane (Lam et al., 2002) (Table 1.1). The activation and translocation of PKC-δ down-regulate the action of insulin by phosphorylating serine/threonine residues on insulin receptor or IRS, thus reducing insulin signaling to downstream targets (Farese, 2001).
In the normal state, approximately 50% of insulin secreted by β-cells is eliminated
by liver during its first pass through the liver (Lewis et al., 2002). Increasing FFA
concentration during the physiological range of portal FFA decreases hepatic insulin
extraction in the perfused rat liver and extraction of insulin is inversely proportional to
hepatic triglyceride contents (Svedberg et al., 1991). Elevated FFA may impair hepatic
extraction of insulin by influencing insulin binding to its receptor (Svedberg et al., 1990).
The reduction in insulin extraction in liver also contributes to hyperinsulinemia.
22 1.4.3 Pancreatic β-cells
The secretion of insulin by the pancreatic β-cell is regulated by many nutrients,
neurotransmitters and hormones (Hedeskov, 1980). Glucose is the key nutrient for
regulating insulin secretion, although many other nutrients including free fatty acids, amino acids and keto acids also affect insulin secretion. There are two signal transduction pathways for glucose-stimulated insulin secretion (GSIS). Metabolism of glucose through
glycolysis increases the ATP/ADP ratio and in turn closes ATP-sensitive K+ channels,
leading to depolarization of the cell membrane and opening of voltage-sensitive Ca2+ channels. The resultant Ca2+ influx increases intracellular Ca2+ concentration and
enhances exocytosis of insulin granules. In addition, glucose metabolism prompts the
production of malonyl-CoA which in turn inhibits CPT-1, leading to the accumulation of
LCFA-CoA in the cytosol. Increased cytosolic LCFA-CoA level may directly enhance
exocytosis and thus insulin secretion by β-cells (Prentki et al., 1997).
The interaction of FFA with pancreatic β-cells seems to be complex. It is well
established that an acute increase in plasma FFA concentration stimulates GSIS in animal
studies (Crespin et al., 1973; Warnotte et al., 1994) and in a human study (Pelkonen et al.,
1968). One possible mechanism is that FFA increase the concentration of LCFA-CoA as
dose glucose (Deeney et al., 2000). A chronic increase in plasma FFA concentration
seems to impair insulin secretion (Boden and Shulman, 2002). It was reported that
Intralipid™ infusion had a biphasic effect on insulin secretion in the perfused pancreas
from rat. An initial stimulation of GSIS after 3-hour Intralipid™ infusion was followed
by a significant inhibition after 24-48 hour infusion, suggesting that a chronic increase in
FFA inhibits GSIS (Sako and Grill, 1990). 23 Chronic FFA increase and concomitant lipid accumulation are toxic to β-cell function by causing β-cell apoptosis (Poitout and Robertson, 2002). DNA fragmentation was increased in rat islets exposed to saturated palmitic acid for 4 days, but not in islets exposed to monounsaturated palmitoleic acid (Maedler et al., 2001). β-cell apoptosis was associated with an increase in ceramide, iNOS expression, and NO production in Zucker diabetic fatty (ZDF) rats (Shimabukuro et al., 1997; Shimabukuro et al., 1998). A recent study of human cadavers also indicated that β-cell mass was decreased in humans with type 2 diabetes compared to controls, and increased β-cell apoptosis rather than decreased neogenesis or replication may be the major mechanism (Butler et al., 2003).
1.4.4 Antilipolytic agents in treatment of type 2 diabetes
Due to the critical role of FFA in the development of insulin resistance and type 2 diabetes, it can be speculated that reduction of FFA availability would ameliorate the metabolic abnormalities caused by insulin resistance and type 2 diabetes. Clinical studies have shown that the new class of antidiabetic agent thiazolidinediones (TZD) enhance insulin sensitivity in part by reduction of plasma FFA concentration and redistribution of intracellular lipid from insulin-responsive organs to peripheral adipocytes (Boden et al.,
2003; Carey et al., 2002; Mayerson et al., 2002). The possible mechanism is that TZD selectively increase lipogenesis in adipocytes by activating PPARγ, thus reducing the availability of circulating FFA for skeletal muscle (Matthaei et al., 2000).
Some clinical studies have shown that reduction of plasma FFA concentration by inhibiting lipolysis improves insulin action. The most frequently used antilipolytic agent in clinical trials is acipimox, a long-acting analog of nicotinic acid (Aktories et al., 1980; 24 Fuccella et al., 1980). Acipimox acutely decreased plasma FFA concentration and improved glucose oxidation and nonoxidative glucose disposal by skeletal muscle in type
2 diabetic patients (Vaag et al., 1991). An overnight administration of acipimox
decreased high plasma FFA levels, thus reducing insulin resistance and improving
glucose tolerance in obese nondiabetic subjects and obese type 2 diabetic patients
(Santomauro et al., 1999). A 7-day administration of acipimox reduced plasma FFA
concentration and increased insulin sensitivity in skeletal muscle and liver in nonobese,
normal-glucose-tolerant human subjects with a family history of type 2 diabetes (Bajaj et
al., 2004).
Collectively, these data support use of antilipolytic agents to ameliorate insulin
resistance and type 2 diabetes. The long-term efficacy and adverse effects of antilipolytic
agents need further evaluation. Because adenosine and PGE2 can potentiate insulin action, the analogs of these antilipolytic agents may become potential candidates in future.
1.5 Ginseng and type 2 diabetes
As discussed above, type 2 diabetes has been a major health problem in North
America. Chronic hyperglycemia may cause many of the complications of type 2 diabetes, including cardiovascular disease, nephropathy, retinopathy, and neuropathy, associated with high morbidity and mortality. Good glycemic control is beneficial for the reduction of the risk of long-term complications (Mudaliar, 2004; Setter et al., 2003).
Conventional treatment strategies for type 2 diabetes include modification of lifestyle, such as diet and exercise, and pharmacological interventions. The main categories of antidiabetic agents include sulphonylureas that stimulate insulin secretion, biguanides
25 that reduce hepatic glucose production, α-glucosidase inhibitors that delay digestion and absorption of intestinal carbohydrate, thiazolidinediones (TZD) that improve insulin
action, and insulin (Krentz and Bailey, 2005; Matthaei et al., 2000; Moller, 2001).
Besides the conventional therapies, the use of complementary and alternative medicine (CAM) to treat diabetes is increasing among diabetic patients (Egede et al.,
2002; Yeh et al., 2002). Herbal remedies and dietary supplements are the common CAM therapies used by people with diabetes (Ryan et al., 2001). A recent systematic review of herbs for glycemic control in diabetes suggested that ginseng was one of several herbs to show efficacy in controlled clinical trials (Yeh et al., 2003). Ginseng has been used to treat diseases for over 3000 years in China (Liu and Xiao, 1992). It is one of the most widely recognized and commonly used herbs around world.
1.5.1 Introduction to ginseng
Ginseng (Ginseng, spp.) has many species, including Korean ginseng (Panax ginseng), American ginseng (Panax quinquefolius), Japanese ginseng (Panax, japonicus), and Siberian ginseng (Eleutherococcus senticosus). Ginseng is considered as a tonic in traditional Chinese medicine (TCM) for restoring and enhancing general vitality. The dried root of ginseng is traditionally used as a medicinal. There are two traditional ways to prepare ginseng. Air-dried ginseng root is called white ginseng, and steam-treated ginseng root is called red ginseng (Gillis, 1997). White ginseng and red ginseng have different effects in TCM. White ginseng is more nourishing to “Yin”, whereas red ginseng is more nourishing to “Qi”. “Yin” and “Qi” are regarded as two basic substances in human body according to the theory of TCM. “Yin” means liquid; “Qi” means energy.
26 According to TCM, ginseng was used to treat a disease corresponding to diabetes with
symptoms of excessive thirst, extreme hunger, frequent urination, and weight loss. It is
assumed that ginseng both nourishes “Yin” and “Qi”.
Current research indicates that ginseng and its active components have multiple
pharmacological effects (Attele et al., 1999), including modulation of immune function
(Friedl et al., 2001; Larsen et al., 2004), enhancement of the ability to learn and memory
(Bao et al., 2005; Kurimoto et al., 2004), inhibition of tumor growth (Keum et al., 2003;
Lee et al., 2005), and improvement of glycemic control (Attele et al., 2002; Vuksan et al.,
2000a).
In general, ginseng is well-tolerated and any adverse effects are mild and transient.
Headache, sleeplessness, and gastrointestinal disorders are the most commonly reported
adverse effects which are usually correlated with overdose of ginseng (Coon and Ernst,
2002). A reported “ginseng-abuse syndrome” included hypertension, nervousness, insomnia, skin eruptions, and morning diarrhea (Siegel, 1979).
1.5.2 Ginsenosides
1.5.2.1 Chemistry
Ginsenosides, a group of steroidal saponins, are the major active components of ginseng, and are used as a marker for quality control of commercial ginseng products
(Harkey et al., 2001). Twenty eight kinds of ginsenosides have been identified, including
12 basic ginsenosides and their metabolites. The basic ginsenosides are divided into three classes based on their chemical structures: panaxadiol (Rb1, Rb2, Rc, Rd, Rg3, Rh2), panaxatriol (Re, Rf, Rg1, Rg2, Rh1), and oleanolic acid (Ro) (Helms, 2004; Liu and Xiao,
27 1992). Panaxadiol and panaxatriol have a 4 trans-ring steroid skeleton with different sugar moieties (Harkey et al., 2001). The addition of sugar moiety to ginsenosides is called glycosylation. The chemical structures of ginsenosides are shown in Figure 1.4.
Ginsenosides are found in different parts of ginseng, including root, leaf, and berry
(Xie et al., 2004a). The concentration of ginsenosides in extracts varies with ginseng species, part of ginseng plant, years of growth, and extraction method (Li et al., 1996).
Ginsenosides Rb1, Rb2, Rc, Rd, Re, Rf and Rg1 are abundant in ginseng (Ji et al., 2001).
Korean ginseng has a higher ratio of Rg1 to Rb1, while American ginseng has a lower ratio of Rg1 to Rb1 (Wang et al., 1999).
1.5.2.2 Metabolism and bioavailability
Accumulating evidence strongly suggests that ginsenosides are prodrugs that are metabolized to their active forms in the human body (Akao et al., 1998; Bae et al., 2002;
Hasegawa et al., 2000; Tawab et al., 2003). Oral bioavailability of intact ginsenosides is very low. The major ginsenosides Rb1 and Rg1 were absorbed from the digestive tract of rats with bioavailability being 4.35% and 18.4% of the dose, respectively (Xu et al.,
2003). Orally administered ginsenoside Rb1 was mostly recovered from the intestine in germ-free rats, suggesting that microbial hydrolysis was required for absorption. The identification of compound K, a metabolite of Rb1, in plasma and in the lower part of intestine of conventional rats supported the need for transformation and absorption (Akao et al., 1998). Thus, ginsenosides are deglycosylated by intestinal bacteria to their active metabolites in the large intestine, and then enter the circulation. Intestinal bacteria cleave sugar moieties stepwise to yield the major metabolites, including 20S-protopanaxadiol
28 20-O-β-D-glucopyranoside (M1) and 20S-protopanaxatriol (M4). After being absorbed from intestine, these metabolites are selectively accumulated in the liver and excreted in bile. Some of these metabolites are further esterified with fatty acids in liver. Esterified metabolites are not excreted as bile and have longer half-life in liver (Hasegawa, 2004;
Hasegawa et al., 2000).
A pilot human study with two subjects focused on the identification of ginsenosides and their hydrolytic products in circulation after oral administration of ginseng extract.
Mass spectrometry revealved that Rh1 and F1, two hydrolysis products of panaxatriol ginsenosides, and compound-K were present in plasma and urine. Very low concentrations of some intact ginsenosides in ginseng extract were detected in urine but not in plasma. This study indicates that bacterial hydrolysis in the gastrointestinal tract plays a key role in ginsenoside bioavailability in humans (Tawab et al., 2003).
1.5.3 Hypoglycemic effect of ginseng
1.5.3.1 Animal studies
Previous studies have shown that extracts from ginseng roots decreased blood glucose level in alloxan-induced diabetic mice (Kimura et al., 1981a) and in streptozotocin-induced diabetic rats (Yokozawa et al., 1985). In recent years, the Yuan group reported a series of studies on hypoglycemic effect of ginseng in ob/ob mice
(genetic deficiency of leptin) and db/db mice (genetic deficiency of the leptin receptor)
(Attele et al., 2002; Dey et al., 2003; Xie et al., 2004a; Xie et al., 2004b; Xie et al., 2002).
Daily intraperitoneal injection of Korean ginseng berry extract (extracted in 75% ethanol) for 12 days decreased fasting blood glucose to normal in obese diabetic mice (C57BL/6J
29 ob/ob). This treatment also was associated with improved glucose tolerance and a significant decrease in fasting and fed serum insulin. The rate of insulin-stimulated glucose disposal measured by hyperinsulinemic-euglycemic clamp increased over two-fold. Weight and food intake of obese diabetic mice were reduced, while energy expenditure was increased. Ginsenoside Re in Korean ginseng berry extract is considered the major active component for this hypoglycemic action (Attele et al., 2002).
Another fraction of American ginseng berry extract that is rich in polysaccharides also has a hypoglycemic effect in ob/ob mice. Daily intraperitoneal injection of ginseng-derived polysaccharides at 150 mg/kg for 10 days decreased fasting blood glucose to normoglycemia compared to the vehicle group. Moreover, there was an extended hypoglycemic effect on the fasting blood glucose level which stayed lower and returned to the control concentration 30 days after terminating treatment with the ginseng-derived polysaccharide. Body weight of ob/ob mice was not affected compared to the vehicle group (Xie et al., 2004b).
1.5.3.2 Clinical studies
Several clinical trials have verified the hypoglycemic effect of ginseng. In a double-blind placebo-controlled study, the effects of ginseng (100 or 200 mg per day, unspecified species) for 8 weeks were studied in 36 newly diagnosed type 2 diabetic patients. The results showed that ginseng therapy reduced fasting blood glucose and body weight and improved mood and psychophysical performance. The 200-mg dose of ginseng also improved A1c hemoglobin (Sotaniemi et al., 1995).
30 The Vuksan group reported a series of short-term and long-term studies on the
hypoglycemic effect of ginseng. Gound root of American ginseng (3 g) taken orally
decreased postprandial glycemia in response to a 25-g oral glucose load in healthy
subjects and subjects with type 2 diabetes (Vuksan et al., 2000a). The hypoglycemic effect did not vary with ginseng dose and time of administration for type 2 diabetic patients. Ginseng doses of 3, 6, or 9 g had similar efficacy, and there was no difference when administered 120, 80, 40, or 0 min before the oral glucose load (Vuksan et al.,
2000b). In contrast, the hypoglycemic effect was time-dependent but not dose-dependent in healthy people. This effect only occurred when the ginseng was administered 40 min before the oral glucose load. Ginseng doses of 1, 2, or 3 g had equivalent hypoglycemic effect (Vuksan et al., 2001).
Two long-term studies both adopted a double-blind, randomized, placebo-controlled, crossover design to evaluate the hypoglycemic effects of American ginseng and Korean red ginseng, respectively. The type 2 diabetic patients in these two studies received ginseng and control treatments while maintaining their conventional diabetes
medications. Administration of American ginseng extract (3 g per day) for 4 weeks
decreased fasting blood glucose and A1c hemoglobin in type 2 diabetic patients (Vuksan
et al., 2000c). Treatment with Korean red ginseng extract (6 g per day) for 6 weeks
improved postprandial glycemia and insulin sensitivity in type 2 diabetic patients
(Vuksan et al., 2003).
31 1.5.4 Mechanisms for the hypoglycemic effect of ginseng
1.5.4.1 Stimulation of cellular glucose uptake
The effects of Panax ginseng extract and ginsenosides on glucose uptake in sheep
erythrocytes were investigated using 2-deoxy-D-[2-3H] glucose (2-DG). Ginseng extract
(extracted in 50% ethanol) at 100 µg/ml increased glucose uptake by 11% compared to
basal level. Most ginsenosides (Rc, Rb2, Rb1, Rf, Rg2, Rg1, and Re) increased 2-DG
uptake by 15-30%. However, ginsenoside Rg3 inhibited 2-DG uptake slightly in sheep
erythrocytes (Hasegawa et al., 1994). The previous work in our lab also showed that
Korean ginseng extract slightly stimulated glucose transport into rat adipocytes in vitro
(Edens et al., 2001).
1.5.4.2 Stimulation of insulin biosynthesis and release
DPG-3-2, a water extract of ginseng, exerted a hypoglycemic effect in
alloxan-induced diabetic mice (Kimura et al., 1981b). A subsequent study (Waki et al.,
1982) showed that DPG-3-2 stimulated insulin biosynthesis in perfused pancreas isolated
from alloxan-induced diabetic rats and genetically diabetic KK-CAy mice. Stimulation of
insulin biosynthesis was determined by incorporation of radioactive leucine into insulin
fractions during a 2-h perfusion of rat pancreas and a 3-h incubation of mouse islets.
DPG-3-2 increased incorporation into insulin 1.5 to 1.8-fold in perfused pancreas from
diabetic but not normal rats. DPG-3-2 also increased insulin biosynthesis in islets from
KK-CAy mice. Moreover, DPG-3-2 promoted insulin release from perfused pancreas of
both normal and diabetic rats (Kimura et al., 1981a).
32 1.5.4.3 Inhibition of glucose absorption from the small intestine
The effects of ginseng root on glucose and maltose transport were investigated in rat and human duodenal mucosa by using short-circuit current (Isc) measurement. In both models, Isc was increased after adding 10 mM glucose or maltose to the mucosal side chamber. This increase in Isc was suppressed by the addition of ground ginseng root (1 g) to the serosal side chamber. This inhibition occurred in a few minutes and was maintained for 1 hour. The results suggest that inhibition of glucose absorption from the small intestine may be one mechanism for the hypoglycemic effect of ginseng (Onomura et al., 1999).
1.5.4.4 Inhibition of lipolysis
Preparations of Korean red ginseng contain adenosine and pyro-glutamic acid.
Korean red ginseng powder inhibited epinephrine-induced lipolysis and stimulated lipogenesis from glucose in the presence of insulin in rat adipocytes. Pyro-glutamic acid both inhibited epinephrine-induced lipolysis and stimulated lipogenesis from glucose in
the presence and absence of insulin (Takaku et al., 1990). EPG-3-2, the methanol extract of ginseng, inhibited isoproterenol-induced lipolysis in epididymal fat pads (Kimura et al.,
1981b).
1.5.5 Summary
Animal and clinical studies have demonstrated that ginseng has a hypoglycemic effect and can ameliorate metabolic abnormalities in type 2 diabetes. The proposed mechanisms include stimulation of cellular glucose uptake, inhibition of glucose
33 absorption from the small intestine, stimulation of insulin biosynthesis and release, and inhibition of lipolysis. The possible active components are ginsenoside Re, panaxans A,
B, C, D and E (glycans), polysaccharides, DPG-3-2 (the water extract), EPG-3-2 (the methanol extract), adenosine, and pyro-glutamic acid. The majority of studies on the mechanisms of ginseng-hypoglycemic effect were published in 1980s or 1990s. In-depth biochemical and molecular mechanisms for the hypoglycemic effect of ginseng are required.
34
Insulin
Insulin Receptor α
β CAP IRS Cbl Grb2 SOS C3G Crk TC10 ras PI3-K
PDK-1 raf
PKCζ/λ PKB MEK mTOR p70S6K GSK PDE3B ERK
Glucose Glycogen Protein Gene Transport↑ Synthesis↑ Lipolysis ↓ Synthesis↑ Transcription
Figure 1.1. Schematic representation of insulin signaling. Insulin binding to its receptor activates the IRS proteins and multiple downstream targets, resulting in diverse physiological effects of insulin.
35
Catecholamines PGE2 Adenosine Insulin
β-AR α2-AR EP3 A1-AR IR
AC Gs Gi PDE3B PI3-K
IRS ATP cAMP AMP PKB
PKA
TG HSL
FFA + Glycerol
Figure 1.2. Schematic representation of regulation of lipolysis. Detailed information can be found in text. β-AR, β-adrenergic receptor; α2-AR, α2-adrenergic receptor; A1-AR, A1 adenosine receptor; IR, insulin receptor; IRS, insulin receptor substrate proteins; AC, adenylate cyclase; Gs, stimulatory GTP-binding protein; Gi, inhibitory GTP-binding protein; PI3-K, phosphatidylinositol 3–kinase; PKB, protein kinase B; PDE3B, phosphodiesterase 3B; PKA, cAMP-dependent protein kinase A; HSL, hormone-sensitive lipase; TG, triglyceride; FFA, free fatty acids.
36
Isoforms Regulation Function
classical PKC Ca2+, phosphatidylserine, inhibition of insulin (cPKCα, β, γ) diacylglycerols (DAG) signaling
novel PKC phosphatidylserine and DAG inhibition of insulin (nPKCδ, ε, η, θ, µ) signaling
atypical PKC phosphatidylserine translocation of GLUT4 (aPKCζ, λ)
Table 1.1. The regulation and function of PKC isoforms in insulin signaling (Jiang and Zhang, 2002).
37
Figure 1.3. Biosynthesis of prostagladins (Narumiya et al., 1999). Conversion of arachidonic acid to PGG2 and then to PGH2 are catalyzed by cyclooxygenase, and sequential conversions of PGH2 to PGD2, PGE2, PGF2α, PGI2, and TXA2 are catalyzed by respective synthase as shown.
38
Continued
Figure 1.4. The chemical structures of ginsenosides (Attele et al., 1999). A, panaxadiols; B, panaxatriols; C, ginsenoside Ro, a nonsteroidal saponin.
39
40
CHAPTER 2
GENERAL METHODS
2.1 Chemicals
Collagenase (Type I) was purchased from Worthington Biomedical (Lakewood, NJ).
Recombinant human insulin (Humulin-R) was purchased from Eli Lilly (Indianapolis,
IN). Bovine serum albumin (BSA, Type V) and adenosine deaminase were purchased from Roche Biochemical (Indianapolis, IN). Isoproterenol (ISO), prostaglandin E2
6 (PGE2), adenosine, N -(2-phenylisopropyl)adenosine (PIA), cilostamide, rolipram, and
other chemicals were purchased from Sigma-Aldrich (St. Louis, MO). Dulbecco’s
modified Eagles’ medium (DMEM) and phosphate-buffered saline (PBS) were purchased
3 from GIBCO (Carlsbad, Ca). [ H] cAMP was purchased from Amersham (Piscataway,
NJ).
Korean ginseng extract (Panax ginseng; KGE; extracted in 50% ethanol) was
obtained from Flachsmann Company (Vista, CA). American ginseng extract (Panax
quinquefolius; AGE; extracted in 80% ethanol) was obtained from Chai-Na-Ta, Inc
(Vancouver, B.C., Canada). Korean ginseng extract was dissolved in propylene glycol
(100 mg/ml) and American ginseng extract was dissolved in water (100 mg/ml) and
stored at –20 ºC until use.
41 2.2 Animal
Young male Sprague-Dawley rats weighing 150 ± 10g were purchased from Harlan
(Indianapolis, IN) and housed in a temperature-and humidity-controlled environment
(12-h light/dark cycle) with free access to tap water and Harlan Teklad 8640 rodent diet.
Rats were acclimated to the laboratory for at least 6 days before the experiment. For lipolysis assay and PDE activity assay, fed rats were killed by decapitation, whereas rats were killed by carbon dioxide inhalation for RNA extraction and Western blot. All animal protocols were reviewed and approved by The Ohio State University Institution
Laboratory Animal Care and Use Committee.
2.3 Adipocyte isolation
The isolation method is a modification of method described by Rodbell (Rodbell,
1964). Approximate 3 g of epididymal and retroperitoneal adipose tissue was removed from rats and placed in the DMEM at 37 ºC. The adipose tissue was minced into small pieces and rinsed three times with PBS at 37 ºC. After weighing, the adipose tissue was incubated in Krebs-Ringer’s-HEPES (KRH) medium with 2.5% BSA, 200 nM adenosine,
5 mM glucose, and 1 mg/ml collagenase at 37°C for 45 min with 80 rpm shaking. When digestion was completed, the digested tissue was filtered with 250 µm nylon mesh.
Isolated adipocytes were washed three times and suspended at a 20% concentration by weighing isolated adipocytes in KRH medium containing 2.5% BSA, 200 nM adenosine, and 5 mM glucose.
42 2.4 Lipolysis assay
200 µl of adipocytes (20% cell suspension) was aliquoted into 2ml of KRH medium
containing 2.5% BSA, 0.8 U/ml adenosine deaminase, and 10 mM glucose. Adipocytes
were incubated with insulin, KGE, PGE2, PIA and PDE inhibitors (cilostamide and
rolipram) at 37°C for 1 hour with shaking at 80 rpm. Glycerol release into incubation
medium was assessed with the Trinder kit (Sigma-Aldrich; St. Louis, MO). The Trinder
kit measures free glycerol using coupled enzymatic reactions. Glycerol is converted to glycerol-1-phosphate (G-1-P) catalyzed by glycerol kinase (GK) in the presence of adenosine-5'-triphosphate (ATP). G-1-P is then oxidized by glycerol phosphate oxidase
(GPO) to dihydroxyacetone phosphate (DAP) and hydrogen peroxide (H2O2). Peroxidase
(POD) catalyzes the coupling of H2O2 with 4-aminoantipyrine (4-AAP) and sodium
N-ethyl-N-(3-sulfopropyl) m-anisidine (ESPA) to produce a quinoneimine dye that
demonstrates a maximal absorbance at 540 nm. The increase in absorbance at 540 nm is
directly proportional to the free glycerol concentration of the sample. Glycerol assay
enzymatic reactions are summarized as below.
GK Glycerol + ATP G-1-P + ADP
GPO G-1-P + O2 DAP + H2O2
POD H2O2 + 4-AAP +ESPA Quinoneimine Dye + H2O2
43 2.5 Adipocyte RNA isolation
Total RNA was extracted from freshly isolated rat adipocytes by the modified
TRIzol method (Invitrogen; Carlsbad, CA). In brief, frozen 400 µl-500 µl packed adipocytes were thawed in 1ml of TRIzol reagent and homogenized by 20 strokes using a
2 ml homogenizer (Wheaton; Millville, NJ). Homogenates were centrifuged at 12,000 × g for 10 min at 4°C, and then the top lipid layer was discarded. 0.2 ml of chloroform was added to extract RNA and the mixture was centrifuged at 12,000 × g for 15 minutes at
4°C. After centrifugation, the colorless upper aqueous phase was transferred to a fresh sterile tube, and 0.5 ml of isopropanol was added to precipitate the RNA at -20 ºC overnight. RNA precipitate was obtained after centrifugation at 12,000 × g for 30 min at
4ºC. Quantity and purity of adipocyte RNA were determined by measuring absorbance at
260 nm and 280 nm by UV-Vis spectroscopy. Integrity of adipocyte RNA was determined by electrophoresis on 1% agarose gel.
2.6 Reverse transcription-polymerase chain reaction (RT-PCR)
RNA (3 µg) was reverse-transcribed using SuperScript First-Strand Synthesis
System for RT-PCR (Invitrogen; Carlsbad, CA). cDNA was amplified by PCR using Taq
DNA Polymerase (Promega; Madison, WI). The published primers (Table 4.2) for
PDE3B (Harndahl et al., 2002) and the four subtypes of PDE4 (Kostic et al., 1997) were used in the present study. All oligonucleotides for primers were synthesized by Integrated
DNA Technologies (Coralville, IA). 2 µl of the first strand cDNA was used as template in
50 µl of reaction buffer. The PCR reactions were carried out by 35 cycles according to
the following protocol: 44 Cycle 1: (1X) 95 °C for 2 min to denature.
Cycle 2: (35X) Step 1 (denaturation): 95 °C for 40 sec
Step 2 (annealing): 55 °C for 30 sec*
Step 3 (extension): 72 °C for 1 min
Cycle 3: (1X) 72 °C for 20 min
Cycle 4: (1X) 4 °C
* The annealing temperature was optimized for each amplified gene, i.e., PDE3B 52 °C,
PDE4A 59 °C, PDE4B 58 °C, PDE4C 58 °C, and PDE4D 59 °C.
The PCR products were resolved by electrophoresis on 1% agarose gel in the
presence of ethidium bromide and visualized by ultraviolet fluorescence. PCR products
were purified by using QIAquick Gel Extraction kit (Qiagene; Santa Clarita, CA) and
sequenced at the DNA Sequencing Facility of The Ohio State University
Neurobiotechnology Center using ABI PRISM BigDye Terminator Cycle Sequencing
Ready Reaction Kits and an ABI 373XL Stretch DNA sequencer (Applied Biosystems;
Foster City, CA).
2.7 Real-time quantitative reverse transcription polymerase chain reaction (QPCR)
Total RNA samples, prepared as discussed in Section 2.5, were quantitated by
UV-Vis spectroscopy. Sample integrity was analyzed using an Agilent 2100 Bioanalyzer
and Eukaryote Total RNA Nano kit (Agilent Technologies; Palo Alto, CA). The primers
and probes (Table 4.8) were designed using the Primer Express software program
(Applied Biosystems; Foster City, CA) to a region of high similarity among PDE3B and
PDE4 (A, B, C, D) sequences (Table 4.9). The TaqMan probes carried a 5’-FAM reporter
45 and a 3’-TAMRA quencher. All oligonucleotides were synthesized by Integrated DNA
Technologies (Coralville, IA). The assay was carried out on DNase I treated samples
using the Platinum Quantitative RT-PCR ThermoScript One-Step kit (Invitrogen;
Carlsbad, CA). The fluorescence intensity of the reporter label was normalized to Texas
Red, the passive reference label added to the buffer. All reactions were performed in an
ABI-PRISM 7900HT Sequence Detection System (Applied Biosystems; Foster City, CA).
Serially diluted samples of Universal Rat Reference RNA (Stratagene; La Jolla, CA) were used to generate a calibration curve for each gene. Each RT-PCR sample was amplified in triplicate. Relative expression levels were determined by the Relative
Standard Curve Method (Applied Biosystems; Foster City, CA). The level of 28s rat RNA
was assayed as an endogenous control for each sample on every reaction plate. The
principle for the Taqman method is shown in Figure 2.1 (Bustin, 2000).
2.8 Western blot analysis
2.8.1 Preparation of crude subcellular fractions
The isolated adipocytes (2 ml of 20% cell suspension in KRH-BSA) were washed
twice with 5 ml of room temperature homogenization buffer containing 40 mM Hepes
(pH 7.4), 10 mM NaF, 1 mM phenylmethylsulfonylfluoride, 0.25 mM sodium orthovanadate, 10 µg/ml antipain, 10 µg/ml leupeptin and 1 µg/ml pepstatin A.
Adipocytes were centrifuged at 125 × g for 2 min at room temperature, resuspended in
0.8 ml of homogenization buffer, homogenized at room temperature with 10 strokes in a
2 ml homogenizer (Wheaton; Millville, NJ) and placed on ice immediately. For cell lysates, homogenates were centrifuged at 1,000 × g for 10 min at 4 °C, and then the fat
46 layer was removed, and the pellets were resuspended. For particulate fraction and cytosolic fraction, homogenates were centrifuged at 16,100 × g for 60 min at 4°C. The fat layer was removed, the infranatant (referred to as cytosolic fraction) was withdrawn, and
the pellet (referred to as particulate fraction) was resuspended in 120 µl of
homogenization buffer. Protein in each fraction was measured with BCA kit using bovine
serum albumin as standard (Pierce; Rockford, IL).
2.8.2 PKB phophorylation
Protein (10 µg) was separated by 7.5% SDS-polyacrylamide gel (SDS-PAGE).
Following electrophoresis, proteins were transferred to nitrocellulose membranes, and
then the membranes were blocked with 5% nonfat milk in PBS plus 0.5 % Tween-20
(PBS-T) overnight at 4 °C. The blot was incubated with phospho-PKB (Ser 473) polyclonal antibody and PKB polyclonal antibodies (Cell Signaling; Beverly, MA) at
1:1000 dilution individually for 2 hrs at room temperature and then washed five times with PBS-T. The washed blot was incubated with horseradish peroxidase conjugated goat anti-rabbit IgG (Cell Signaling; Beverly, MA) at 1:2000 dilution for 1 hr at room temperature, then washed five times with PBS-T. Bands were detected using the
Supersignal chemiluminescence (Pierce; Rockford, IL).
2.8.3 PDE3B and PDE4B expression
Protein (20 µg) was separated by 7.5% SDS-PAGE. Following electrophoresis, proteins were transferred to nitrocellulose membranes, and then the membranes were blocked with 5% nonfat milk in PBS-T for 2 hrs at room temperature. The blot was 47 incubated with rat PDE3B N-terminal antibody (provided by Vincent Manganiello) at
1:1000 dilution and PDE4B C-terminal antibody (FabGennix; Shreveport, LA) at 1:500
overnight at 4 °C, and then washed five times with PBS-T. The washed blot was
incubated with horseradish peroxidase conjugated goat anti-rabbit IgG (Cell Signaling;
Beverly, MA) at 1:4000 dilution for 1 hr at room temperature, then washed five times
with PBS-T. Bands were detected using the Supersignal chemiluminescence (Pierce;
Rockford, IL). For the negative controls, the blot was incubated with normal rabbit IgG
(Upstate; Lake Placid, NY) at 1:2000 dilution overnight at 4 °C, and then incubated with
horseradish peroxidase conjugated goat anti-rabbit IgG (Cell Signaling; Beverly, MA) at
1:4000 dilution for 1 hr at room temperature.
2.9 Purification of [3H] cAMP
2.9.1 Thin layer chromatography (TLC) purification
250 µl of 1 mCi/ml [3H] cAMP (Amersham; Piscataway, NJ) was applied in 0.5 µl
spots on a line 2 cm above the bottom edge of TLC cellulous fluorescent plate (EMD
Chemicals; Gibbstown, NJ), dried with a cool air stream. As controls, 10 µl of cold
cAMP (20 mM) was applied on either side of the [3H] material. The TLC plate was
placed in TLC tank filled with 50 ml of TLC buffer (5:2 alcohol : 0.5 M ammonium acetate), run for 4 hours, and then dried. Control spots were identified with UV lamp. The corresponding rectangular area that contains [3H] cAMP band was scraped out, and resin
was collected and transferred to a 15 ml centrifuge tube. The [3H] cAMP was eluted with
2 ml of water four times. The pooled eluate was stored at -20 °C in 1 ml aliquots.
48 2.9.2 Column purification
1 ml of TLC-purified [3H] cAMP was applied to the washed DEAE-Sephadex A-25 column, and the flowthrough was discarded. After washing the column with 10 ml of
water two times, the column was eluted with 10 ml of 50 mM HCl. The eluate was
collected in 1 ml fractions and counted. The highest count fractions were pooled to
achieve around 30,000 cpm/10 µl, then were mixed with 0.3 µM cAMP (cold) in PDE
assay buffer containing 50 mM HEPES, pH 7.4, 0.1 mM EGTA, and 8.3 mM MgCl2, with the final count 25,000 to 35,000 cpm/100 µl. The substrate solution was stored at
-20 °C in 2 ml aliquots.
2.10 Phosphodiesterase activity assay
Isolated adipocytes (2 ml of 20% cell suspension in KRH-BSA) were washed twice with 5ml of homogenization buffer containing 50 mM Tris (pH 7.4), 250 mM sucrose, 1 mM EDTA, 0.1mM EGTA, 10 µg/ml antipain, 10 µg/ml leupeptin, 1 µg/ml pepstatin A.
Adipocytes were centrifuged at 125 × g for 2 min at room temperature, resuspended in
0.8 ml of homogenization buffer with the addition of 100 nM calyculin A, homogenized at room temperature with 10 strokes in a 2 ml homogenizer (Wheaton; Millville, NJ), and then placed on ice immediately.
Subcellular fractions were prepared for PDE activity assay according to the method described by Manganiello (Manganiello and Vaughan, 1973). (1) For the subcellular PDE activity distribution, defatted homogenates (H) were centrifuged at 10,000 × g for 7 min at 4 °C and the 10,000 × g pellet fraction was referred to as P1. The supernatant was then centrifuged at 100,000 × g for 20 min and the 100,000 × g pellet fraction was referred to 49 as P2. Each fraction pellet was resuspended in 200 µl of homogenization buffer. The
100,000 × g supernatant fraction was referred to as S. (2) For multiple incubations, the whole particulate fraction was needed only. Defatted homogenates were directly centrifuged at 100,000 × g for 20 min at 4 °C and this 100,000 × g pellet fraction was referred to as particulate fraction. The pellet was resuspended in 200 µl of homogenization buffer. Protein in each fraction was measured with BCA kit using bovine serum albumin as standard (Pierce; Rockford, IL).
PDE activity was determined as previously described by Manganiello (Manganiello and Vaughan, 1973). Samples were incubated at 30°C for 10 min in a total volume of 300
3 µl containing 50 mM HEPES, pH 7.4, 0.1 mM EGTA, 8.3 mM MgCl2 and 0.1 µM [ H] cAMP (25-35,000 cpm) as substrate. After incubation with Crotalus atrox venom at 30
°C for 30 min, 5’-AMP was dephosphorylated to adenosine. Adenosine was separated from substrate using DEAE-Sephadex A-25 columns and quantified by scintillation counting. Samples were diluted so that hydrolysis of substrate was less than 20%. PDE activity is expressed as the amount of cAMP converted to AMP per minute per mg of protein. The activity of each PDE isoform was calculated by the difference in activity in the absence and presence of a specific isoform inhibitor.
2.11 Statistical analysis
Data are presented as mean ± SEM. For the lipolysis assay using cilostamide and rolipram, two-way ANOVA (SPSS 12.0; Chicago, IL) was performed to determine whether treatment and inhibitor had a significant effect, respectively. When a significant effect was found, a priori Bonferroni t tests were used to compare 8 pairs of interests 50 between each treatment group (insulin, KGE, PGE2, and PIA) and basal group. The α level was set at 0.05/8=0.006. No subtle comparison among each group was made.
Student’s t test was used to evaluate statistical significance of the synergistic effect of combining PDE3 and PDE4 inhibitors, and the α level was set at 0.05. A synergistic effect was defined as greater than the sum of individual effects. For basal lipolysis assay and PDE activity assay, data were analyzed by Student’s t test and the α level was set at
0.05.
51
Figure 2.1. The principle for the Taqman method of QPCR (Bustin, 2000). (A) cDNA is obtained from the reverse-transcription of mRNA. (B) Primers and probe are hybridized to their target sequences after denaturation. The fluorescent signal from the probe is quenched because the fluorescent dye and the quenching dye are close. (C) cDNA is amplified by Taq DNA Polymerase. (D) The 5’-nuclease activity of polymerase cleaves the probe. (E) The fluorescent dye and the quenching dye are separated, and the fluorescent signal is detected. The intensity of fluorescence increases in each cycle, directly proportional to the amount of the cleaved probe. 52
CHAPTER 3
MECHANISMS FOR GINSENG ANTILIPOLYSIS IN RAT ADIPOCYTES
IN VITRO
3.1 Introduction
The antilipolytic effect of ginseng has been reported previously (Kimura et al.,
1981a; Takaku et al., 1990). It was thought that ginseng contains insulin-like constituents
(Ng and Yeung, 1985). However, the mechanisms underlying the antilipolytic effect of
ginseng remain unclear. A recent study in our laboratory compared the antilipolytic effect
of American ginseng and insulin (Reaves and Edens, 2001). This study showed that
American ginseng extract (AGE; Chai-Na-Ta; Vancouver, B.C., Canada) mimicked the
antilipolytic effect of insulin in isolated rat adipocytes. AGE inhibited lipolysis in a
dose-dependent manner with ED50 at 63 µg/ml. Like insulin, AGE significantly inhibited
basal lipolysis but not isoproterenol (3 µM)-stimulated lipolysis, suggesting that the effect of AGE is not due to nonspecific damage to adipocytes. Six major ginsenosides
(Rg1, Re, Rb1, Rc, Rb2, and Rd) were detected in AGE by HPLC, but individual ginsenosides were not responsible for AGE antilipolysis.
It was also found in this study that AGE and insulin inhibited both glycerol and FFA release from rat adipocytes. However, the FFA:glycerol ratio for insulin ranged from 3.1
53 to 0.4, and the FFA:glycerol ratio for AGE varied only from 2.3 to 2.1. This difference
may result from the different pattern of FFA re-esterification between insulin and AGE.
Unlike glycerol release, the net release of FFA from adipocytes is determined by two
processes, lipolysis and FFA re-esterification. It is known that insulin has significant
effects on both lipolysis and FFA re-esterification (Campbell et al., 1992; Van Harmelen
et al., 1999). Increasing the concentration of insulin should both reduce lipolysis and
increase FFA re-esterification, thus decreasing the FFA:glycerol ratio in a
concentration-dependent manner. In contrast, the FFA:glycerol ratio for AGE was
independent of concentration, suggesting that AGE reduced lipolysis but did not increase
FFA re-esterification. In contrast, it was reported in the previous study that pyro-glutamic
acid, an antilipolytic component of Korean red ginseng, slightly stimulated lipogenesis
from glucose in the presence and absence of insulin in rat adipocytes (Takaku et al.,
1990).
The molecular mechanisms by which AGE inhibits lipolysis were investigated in
this study. The PI3-K inhibitor, wortmannin (100 nM), partially reduced the antilipolytic
effect of insulin, but did not affect the antilipolytic effect of AGE. The nonselective PDE
inhibitor, enprofylline (1 mM), completely reversed the antilipolytic effects of both
insulin and AGE. The specific PDE3 inhibitor, cilostamide (5 µM), completely reversed
insulin antilipolysis, but partially reduced AGE antilipolysis. The results of this study
suggested that AGE antilipolysis was not mediated by PI3-K, and was dependent on
phosphodiesterase, but not entirely dependent on PDE3. A subsequent study in our lab
showed that the PDE4 specific inhibitor rolipram reduced ginseng antilipolysis by 20%,
54 but not insulin antilipolysis (Reaves and Edens, 2002). Taken together, it can be inferred
that there is divergence in signal transduction for ginseng and insulin.
The purpose of the present study was to investigate if KGE antilipolysis is mediated by the combination of PDE3 and PDE4 and through activation of PKB independent of
PI3-K in rat adipocytes.
3.2 Effects of insulin and KGE on lipolysis
Rationale: The PDE3 inhibitor cilostamide and the PDE4 inhibitor rolipram are
specific PDE isoform inhibitors. When incubating isolated adipocytes with the PDE3 and
PDE4 inhibitors, the effects mediated by these two enzymes are removed. If both PDE3
and PDE4 are activated in ginseng antilipolysis, then inhibition of these two PDE
isoforms should reverse ginseng lipolysis.
Hypothesis: If ginseng antilipolysis is dependent on PDE and is not completely reversed by the PDE3 and PDE4 inhibitors alone, then ginseng antilipolysis is mediated by combination of PDE3 and PDE4.
Experimental design: Freshly isolated adipocytes were incubated with the specific
PDE3 inhibitor cilostamide (5 µM), the specific PDE4 inhibitor rolipram (10 µM), and the combination of cilostamide (5 µM) and rolipram (10 µM) in the absence and presence of insulin (90 pM) and KGE (100 µg/ml). In preliminary experiments, the antilipolytic effects of AGE and KGE were compared and found to be similar (Figure 3.1). Insulin at
90 pM and KGE at 100 µg/ml both induce a half-maximal inhibition of basal lipolysis; cilostamide at 5 µM (Reaves and Edens, 2001) and rolipram at 10 µM
55 (Reaves and Edens, 2002) are the lowest doses which have maximal effects on reduction
of lipolysis. Detailed information on rat adipocyte isolation and lipolysis assay can be
found in Section 2.3 and 2.4 respectively.
Results: Insulin inhibited lipolysis by 42.4% compared to basal (n=8; p<0.002; Figure
3.2). As expected, the PDE3 inhibitor, cilostamide, completely reversed insulin antilipolysis compared to basal with cilostamide. The PDE4 inhibitor, rolipram, did not
reduce insulin antilipolysis.
KGE inhibited lipolysis by 49% compared to basal (p<0.002). Cilostamide reduced
KGE antilipolysis to 43% compared to basal with cilostamide, and KGE inhibited lipolysis significantly (p<0.002). Rolipram reduced KGE antilipolysis to 23.4%
compared to basal with cilostamide, and KGE still inhibited lipolysis significantly
(p<0.002). The combination of these two inhibitors completely reversed KGE
antilipolysis compared to basal with the two inhibitors.
3.3 Effects of insulin and KGE on PKB phosphorylation and translocation
Rationale: Activation of PDE3B by insulin is dependent on a series of phosphorylation
cascades, including insulin receptor, IRS, PI3-K, and PKB. PKB is the critical step which
links PI3-K and PDE3B. It is known that ginseng antilipolysis is not mediated by PI3-K
(Reaves and Edens, 2001). However, it is possible that PKB is activated by
Ca2+/calmodulin protein kinase pathway independent of PI3-K (Okuno et al., 2000). If
PKB is activated by a PI3-K-independent pathway, then its downstream target PDE3B
and possibly PDE4 can be activated.
56 In the insulin signaling pathway, the generation of PIP3 by PI3-K on the plasma
membrane results in recruitment and phosphorylation of PKB. Translocation of PKB to
plasma membrane was detected with a maximal effect after 7 min in rat adipocytes upon
insulin activation (Göransson et al., 1998). If ginseng activates PKB, then translocation of
PKB to the plasma membrane is detected.
Hypothesis: If ginseng activates PKB through a PI3-K independent pathway, then phosphorylation and translocation of PKB to plasma membrane will be detected.
Experimental design: Freshly isolated rat adipocytes were incubated with insulin (900
pM) and a range of concentrations of KGE (125 µg/ml, 250 µg/ml, 500 µg/ml, and 1000
µg/ml) for 10 minutes. The dose and time course for insulin stimulation in the present study were based on the previous study (Göransson et al., 1998), in which it was shown that PKB phosphorylation stimulated by 1 nM insulin was clearly detected after 7 min.
The dose range for ginseng was based on the antilipolytic effect of ginseng.
Phospho-PKB (Ser 473) antibody and PKB antibody was used to detect phospho-PKB and total PKB, respectively. For the ginseng-dose experiment, the proteins in adipocyte homogenates were subjected to SDS-PAGE and Western blot. For the experiment to distinguish the distribution of phospho-PKB, the proteins in homogenates, the particulate fraction, and the cytosolic fraction were subjected to SDS-PAGE and Western blot.
Detailed information on preparation of adipocyte subcellular fractions and Western blot of PKB phosphorylation can be found in Section 2.8.1 and 2.8.2 respectively.
Results: Contrary to the hypothesis, KGE did not increase PKB phosphorylation
(Figure 3.3). As expected, insulin, the positive control, increased PKB phosphorylation
in defatted cell lysates. The increased PKB phosphorylation was detected in cell lysates, 57 cytosolic fraction, and particulate fraction of adipocytes treated with insulin (Figure 3.4).
The results of total PKB were shown as a control. After 10 min stimulation with insulin,
PKB phosphorylation in the cytosol fraction was still stronger than that in the particulate
fraction. No significant PKB translocation was observed.
3.4 Discussion
The present study aimed to investigate the antilipolytic effect of KGE in rat adipocytes and the intracellular signaling pathway for KGE antilipolysis. It is known that
PDE3B is the key enzyme contributing to the antilipolytic action of insulin in adipocytes.
The present study showed KGE inhibits lipolysis like insulin, but the signaling pathway
for KGE antilipolysis is different from that activated by insulin. PKB activation is not
involved in KGE antilipolysis. Moreover, PDE3B is not the only downstream target for
KGE antilipolysis, and KGE antilipolysis is also mediated by PDE4.
The signaling pathways that mediate KGE and insulin antilipolysis were compared.
For insulin, the signaling pathway includes a cascade of phosphorylation from the insulin
receptor through PDE3B, including IRS, PI3-K, and PKB (Shakur et al., 2001). The data
from the previous study demonstrated that AGE antilipolysis is independent of PI3-K
(Reaves and Edens, 2001). It is known that PKB is activated through not only the PI3-K
pathway but also the Ca2+/calmodulin protein kinase pathway. PKB was reported to be phosphorylated at Thr-308 by Ca2+/calmodulin-dependent protein kinase kinase alpha
(CaM-kinase kinase alpha) (Okuno et al., 2000). Moreover, an increase in intracellular calcium caused by KCl-induced depolarization inhibited lipolysis stimulated by several agonists in human adipocytes (Xue et al., 2001). Therefore, it was initially hypothesized
58 that a ginseng-induced increase in intracellular calcium concentration activates the
Ca2+/calmodulin-dependent protein kinase pathway, and thus PKB is phosphorylated by
CaM-kinase kinase alpha independent of PI3-K. However, the data from the present
study showed that KGE did not affect PKB phosphorylation. Inconsistent with the
previous study (Göransson et al., 1998), PKB translocation stimulated by insulin was not
observed in the present study. A possible reason for this discrepancy was the difference in
centrifugation force (16,100 × g in the present study vs. 33,000 × g in Göransson et al.’s
study) for preparation of membrane fraction.
Whether specific PDE(s) are the downstream target for ginseng was further
determined. Both PDE3 and PDE4 hydrolyze cAMP to AMP (Francis et al., 2001).
Although information on the expression of PDE4 in adipocytes is not available, PDE4
activity has been detected in adipocytes by using biochemical and pharmacological
methods (Schmitz-Peiffer et al., 1992; Zhang and Carey, 2004). Moreover, inhibition of
PDE4 increased basal lipolysis in 3T3-L1 adipocytes, rat adipocytes, and human
adipocytes (Elks and Manganiello, 1984; Nakamura et al., 2004; Snyder et al., 2005; Xue
et al., 2001). However, a PDE4 inhibitor did not eliminate the antilipolytic effect of
insulin (Shechter, 1984). The novel finding in the present study is that the combination of
PDE3 and PDE4 inhibitors completely reversed KGE antilipolysis. Therefore, it is
reasonable to hypothesize that KGE may inhibit lipolysis by activating PDE3 and PDE4
through an alternative signaling pathway.
The reported active components for the antilipolytic effect of ginseng include
EPG-3-2 (the methanol extract), adenosine, and pyro-glutamic acid (Kimura et al., 1981a;
Takaku et al., 1990). EPG-3-2 contained primarily unknown substances and minor
59 concentrations of ginsenosides Rg and Re (Kimura et al., 1981a). Ginsenosides are considered the main active constituents of ginseng. Korean ginseng and American ginseng have distinct ginsenoside profiles. The ratio of ginsenoside Rg1 to Rb1 (Rg1/Rb1) is generally higher in Korean ginseng than in American ginseng (Harkey et al., 2001).
The concentration of six major ginsenosides in AGE and KGE was determined by HPLC method (Table 3.1; Edens, personal communication, 2003). The values of Rg1/Rb1 were
0.21 for KGE and 0.08 for AGE, respectively. Although it was reported that ginsenoside
Re in Korean ginseng berry extract was the major active component for the hypoglycemic action (Attele et al., 2002), no individual ginsenosides including Re inhibit lipolysis (Reaves and Edens, 2001). In addition, the equivalent antilypolytic effects of
KGE and AGE was independent of ginsenoside profiles that differes in the concentrations of ginsenosides Rg1, Rc, Rb2, and Rd. Thus, ginsenosides are not the key antilipolytic components in ginseng extract.
Adenosine and pyro-glutamic acid are present in Korean red ginseng powder
(Takaku et al., 1990). Adenosine is an antilipolytic agent (Liang et al., 2002). Addition of adenosine deaminase to the incubation hydrolyzes adenosine and prevents this antilipolytic effect (Honnor et al., 1985). The concentration of adenosine deaminase (0.8
U/ml) used in the present study was sufficient to eliminate the antilipolytic effect of adenosine in ginseng (Reaves and Edens, 2001). However, KGE and AGE inhibited lipolysis similar to insulin even in the presence of adenosine deaminase in the present study. Therefore, some active components other than adenosine likely contribute to the
60 antilipolytic effect of ginseng. Currently, it remains unknown if the antilipolytic effects of
AGE and KGE in the present study are completely attributable to pyro-glutamic acid.
In summary, the present study shows that KGE, like insulin, inhibits lipolysis in rat
adipocytes and that KGE antilipolysis may be mediated in part by PDE4. Compared with
insulin, there appears to be an alternative signaling pathway for KGE to activate PDE4 in
adipocytes. In addition to PDE3B, PDE4 activity is present in adipocytes and may
contribute to antilipolysis. Therefore, ginseng has the potential to serve as a useful tool to investigate an alternative signaling pathway for activating PDE4 in rat adipocytes.
61
500
M) µ 400
300
200
100
Glycerol Concentration ( Concentration Glycerol 0 Basal Insulin AGE KGE
Figure 3.1. Comparison of AGE and KGE antilipolysis in rat adipocytes. Rat adipocytes were incubated with insulin (150 pM), AGE (100 µg/ml), and KGE (100 µg/ml). Basal indicates that no treatment is present. Data are mean ± SEM of 3 independent experiments assayed in triplicate.
62
CON CIL ROL CIL+ROL
500 mol/L) µ 400 * 300 * * 200 * *
100
0
Glycerol Concentration ( Basal Insulin KGE
Figure 3.2. Effects of insulin and KGE on lipolysis in rat adipocytes. Rat adipocytes were incubated with the specific PDE3 inhibitor cilostamide (CIL, 5 µM), the specific PDE4 inhibitor rolipram (ROL, 10 µM), and combination of cilostamide (5 µM) and rolipram (10 µM) in the absence and presence of insulin (90 pM) and KGE (100 µg/ml). Basal indicates that no insulin or KGE is present. Control (CON) indicates that no PDE inhibitor was present. Data are mean ± SEM of 5 independent experiments assayed in triplicate. Statistical significance compared to basal for each inhibitor is denoted with *, P <0.006.
63
phospho-PKB (59KD)
PKB (59 kDa)
Con KGE KGE KGE KGE Ins 900 125 250 500 1000 pmol/L µg/ml µg/ml µg/ml µg/ml
Figure 3.3. Effects of insulin and a range of concentrations of KGE on PKB phosphorylation in cell lysates of rat adipocytes. Adipocytes were incubated with insulin (Ins, 900 pM) and KGE (125 to 1000 µg/ml) for 10 min. Control (Con) represents sample from adipocytes incubated without insulin or KGE. Cell lysates were prepared as described in Section 2.8.1. Protein (10 µg) was subjected to SDS-PAGE and Western blot using phospho-PKB (Ser 473) and PKB antibody. Results shown were representative of 3 independent experiments. KGE did not affect phosphorylation of PKB, the downstream target of PI3-K, whereas insulin increased phosphorylation of PKB.
64
phospho-PKB (59KD)
PKB (59 kDa)
Con Ins KGE Con Ins KGE Con Ins KGE
Cell lysates Cytosolic Particulate fraction fraction
Figure 3.4. Effects of insulin and KGE on PKB phosphorylation in cell lysates, cytosolic fraction, and particulate fraction of rat adipocytes. Adipocytes were incubated with insulin (Ins, 900 pM) and KGE (500 µg/ml). Control (Con) represents sample from adipocytes incubated without insulin or KGE. Cell lysates, cytosolic fraction, and particulate fraction were prepared as described in Section 2.8.1. Protein (10 µg) was subjected to SDS-PAGE and Western blot using phospho-PKB (Ser 473) and PKB antibody. Results shown were representative of 3 independent experiments. The increased PKB phosphorylation was detected in cell lysates, cytosolic fraction, and particulate fraction of adipocytes incubated with insulin. PKB phosphorylation in the cytosolic fraction was still stronger than that in the particulate fraction. No significant PKB translocation to particulate fraction was observed.
65
Ginseng Rg1 Re Rb1 Rc Rb2 Rd species (g/100 g)
AGE 0.31 2.40 3.83 0.54 0.11 0.84
KGE 0.85 2.74 4.02 2.79 2.91 2.12
Table 3.1. Comparison of ginsenoside profiles between American ginseng extract (AGE) and Korean ginseng extract (KGE). AGE and KGE were analyzed for concentration of six major ginsenosides by HPLC (Edens, personal communication, 2003).
66
CHAPTER 4
THE EXPRESSION OF PHOSPHODIESTERASE 4 IN RAT ADIPOCYTES
4.1 Introduction
Cyclic adenosine monophosphate (cAMP) and cyclic guanosine monophosphate
(cGMP) are important intracellular second messengers and mediate multiple
physiological functions in response to extracellular stimuli (Lincoln, 1989; Maurice et al.,
2003; Murray, 1990). cAMP and cGMP are degraded to AMP and GMP by cyclic nucleotide phosphodiesterase (PDE) (Conti and Jin, 1999). PDE are a large family of structurally related enzymes. To date, at least eleven families of PDEs (PDE1 to 11) have been reported (Beavo and Brunton, 2002). For each PDE family, multiple isoforms are generated as a result of different genes or different splicing variants (Houslay, 2001).
PDEs differ in affinity for substrate (cAMP or/and cGMP), response to specific effectors,
and sensitivity to specific inhibitors (Francis et al., 2001). Among the family of PDEs,
PDE4, PDE7, and PDE8 specifically hydrolyze cAMP. PDE5, PDE6, and PDE9
specifically hydrolyze cGMP. Finally, PDE1, PDE2, PDE3, PDE10, and PDE11
hydrolyze both cAMP and cGMP (Francis et al., 2001).
The general structure of PDE is composed of three regions, VIZ., N-terminal
regulatory domain, catalytic domain, and C-terminal domain (Shakur et al., 2001). The
67 catalytic domains of PDEs are highly conserved for the same gene family. The
N-terminal regulatory domains are variable among PDEs and contain different sites that
regulate PDE catalytic activity in response to specific signals. Binding to divalent cations
(Zn 2+ and Mg 2+) in the catalytic domain is necessary for the catalytic activity of all
PDEs (Francis et al., 2001).
Phosphodiesterase 3 (PDE3) has two gene families, PDE3A and PDE3B. PDE3 has
high affinity for both cAMP and cGMP. Because cGMP inhibits PDE3-mediated cAMP
hydrolysis, it was referred to as cGMP-inhibited PDE in the old nomenclature. The
pharmacological inhibitors for PDE3 include cilostamide, OPC3911, and milrinone.
PDE3A and PDE3B are ~120 kDa and ~135 kDa proteins, respectively. PDE3A exists in
the cytosolic fraction, whereas PDE3B is a membrane-associated protein and its
N-terminal region contains 5-6 transmembrane helices (Shakur et al., 2001).
Expression of PDE3A and PDE3B is tissue-specific. PDE3A is abundant in cardiac
tissue, platelets, and vascular smooth muscle, and PDE3B is abundant in hepatocytes,
adipose tissue, and pancreas. The physiological functions of PDE3 are inferred from
experiments with specific inhibitors of PDE3. PDE3 reduces heart contractility, increases
smooth muscle contraction, promotes platelet aggregation, decreases insulin exocytosis,
and inhibits lipolysis and glycogenolysis (Manganiello et al., 1995; Pyne and Furman,
2003; Shakur et al., 2001).
PDE4 is encoded by four genes (A, B, C, D) and consists of more than16 splice
variants. PDE4 specifically hydrolyzes cAMP and was referred to as cAMP-specific PDE
in the old nomenclature. The pharmacological inhibitors for PDE4 include rolipram and
Ro 20-1724 (Houslay, 2001). PDE4 subtypes exist in brain, heart, liver, lung, kidney,
68 immune cells, blood cells, and preadipocytes (MacKenzie et al., 1998; Muller et al.,
1996). The specific inhibitors of PDE4 have provided some information on the physiological role of PDE4. PDE4 plays a regulatory role in inflammation and memory
(Houslay, 2001). The PDE4 inhibitors inhibit inflammation (Jimenez et al., 2004;
Manning et al., 1999) and enhance memory (Bourtchouladze et al., 2003; Zhang et al.,
2000). The major characteristics of PDE3 and PDE4 are compared in Table 4.1.
PDE4 proteins are located in cytoplasm and membranes (Houslay, 2001). All three human PDE4B isoforms (PDE4B1, PDE4B2, and PDE4B3) were present in both the cytosolic and particulate fractions of transfected COS-7 cells (Huston et al., 1997).
PDE4A1 (Pooley et al., 1997) was exclusively located in the particulate fraction of stably
transfected human thyroid carcinoma FTC cell lines. PDE4A4 (Huston et al., 1996),
PDE4A5 (McPhee et al., 1995), and PDE4A8 (Bolger et al., 1996) were each located in
the particulate and cytosolic fractions of transfected COS-7 cells. PDE4D short isoforms,
PDE4D1 and PDE4D2, were found only in the cytosolic fraction. In contrast, the PDE4D
long isoforms, PDE4D3, PDE4D4, and PDE4D5, existed in both the cytosolic and
particulate fractions of COS-7 cells (Bolger et al., 1997).
PDE4 has two highly conserved regions, called upstream conserved regions 1 and 2
(UCR1 and UCR2) in the N-terminal regulatory domain. The long form of PDE4
contains UCR1 and UCR2; and the short form of PDE4 contains UCR2 or truncated
UCR2. Short-term regulation of PDE4 activity is dependent on phosphorylation of the
protein. PDE4 long isoforms can be activated through PKA phosphorylation at Ser-54 in
UCR1 (Conti et al., 2003; Houslay, 2001).
69 In addition, both long forms and short forms of PDE4 isoforms are differentially
regulated at the catalytic unit through ERK phosphorylation. For PDE4 long isoforms,
ERK phosphorylation has an inhibitory effect on activity. However, this effect can increase cAMP concentrations which activate PKA. Therefore, this feedback regulation generates the net activation of PDE4 long isoforms. In contrast, ERK phosphorylation directly activate PDE4 short isoforms directly (Houslay and Adams, 2003).
PDE4 activity has been detected by biochemical and pharmacological methods in rat adipocytes (Schmitz-Peiffer et al., 1992; Shechter, 1984) and 3T3-L1 adipocytes (Elks and Manganiello, 1984; Zhang and Carey, 2004), but the expression and regulation of
PDE4 in rat adipocytes has not been investigated. The purpose of this study was to determine the gene and protein expression of PDE4 in rat adipocytes.
4.2 Determination of PDE4 gene expression in rat adipocytes
Rationale: Gene expression of PDE4 subtypes has been shown in a variety of organs,
tissue, and cells (Muller et al., 1996). However, there is no available data on gene
expression of PDE4 subtypes in rat adipocytes. It has been demonstrated that PDE4
activity exists in rat adipocytes using biochemical methods. In the ginseng study, PDE4
activity was detected by use of the specific PDE4 inhibitor, rolipram. Therefore, the
PDE4 gene is expected to be expressed in rat adipocytes. In the following experiment,
expression of the PDE4 gene in rat adipocytes will be determined by RT-PCR.
Hypothesis: If specific inhibitors of PDE4 reverse the effects of antilipolytic agents,
PDE4 is expressed in rat adipocytes.
70 Experimental design: Published PDE3B and generic PDE4 primers were used (Table
4.2). The generic PDE4 primers were designed to amplify regions in catalytic units
common to all known splice variants in a specific subtype of PDE4 (Kostic et al., 1997).
It is known that PDE3B mRNA is present in rat adipocytes (Liu and Maurice, 1998), so it
was used as an internal control for this study. Detailed information on adipocyte RNA
isolation and RT-PCR can be found in Section 2.5 and Section 2.6 respectively.
Results: PCR products encoding PDE3B (530bp), PDE4A (233bp), PDE4B (786bp),
PDE4C (539bp), and PDE4D (262bp) sequences were amplified (Figure. 4.1).
Amplification reactions performed using RNA that had not been reverse-transcribed yielded no PCR products. Because the rat adipocyte RNA samples for RT-PCR were not treated with DNase, there was genomic DNA contamination in the samples, leading to nonspecific bands around or over 2000 bp in the reverse-transcribed RNA samples and the nonreverse-transcribed RNA samples of PDE4A and PDE4C. Results similar to the results shown in Figure 4.1 were obtained using RNA extracted from three independent preparations of isolated rat adipocytes. Sequencing of the PCR products of PDE3B and
PDE4 (A, B, C, D) showed that they were identical to the previously published sequences
using NCBI two-sequence BLAST (Table 4.3-4.7).
4.3 Quantification of PDE4 gene expression in rat adipocytes
Rationale: Based on the previous and present studies, both PDE3B and PDE4 are
expressed in rat adipocytes. The previous study showed that the PDE3 activity accounted
for more than 90% of the total PDE activity in the particulate fraction and the PDE4
activity accounted for 10% of the total PDE in the whole rat adipocytes (Eriksson et al.,
71 1995). It is of interest to know the expression of PDE4 gene relative to that of PDE3B gene in rat adipocytes. Moreover, it needs to be clarified which PDE4 subtype is predominant among four PDE4 subtypes in rat adipocytes. In the following experiment, the Taqman method of QPCR (real-time quantitative reverse transcription polymerase chain reaction) was used because this method is more specific than the SYBR green method (Bustin, 2000).
There are two strategies to quantitate gene expression: absolute quatification and relative quantification. Absolute quantification relates the PCR signal to input copy number using an absolute standard curve, while relative quantification measures the levels of target mRNA expression relative to the levels of an internal control RNA.
Relative quantification is easier to perform than absolute quantification and is adequate to serve most investigations of gene expression (Bustin, 2000). In the following experiment, relative quantification was used to measure PDE3B and PDE4 gene expression normalized to 28s rat RNA.
Hypothesis: If the proportion of PDE3 activity exceeds that of PDE4 activity in rat adipocytes, then expression of PDE3B will be higher than that of PDE4.
Experimental design: The primer and probe sets for PDE3B and PDE4 (A, B, C, D) genes (Table 4.8) were designed to a region of homology across the five genes (Table 4.9) to ensure equivalent reverse-transcription efficiency. BLAST analysis of the amplicon sequences did not yield any significant cross-alignments (data not shown). The standard curves for the amplification of PDE3B and PDE4 (A, B, C, D) and three calibration curves for each target gene are shown in Figure 4.2. The primer sets generated PCR amplification efficiencies of 94% for PDE3B, 101% for PDE4A, 96% for PDE4B, 66%
72 for PDE4C, and 91% for PDE4D (Table 4.10). There are two methods to treat data in
relative quantification, comparative CT method and relative standard curve method. The
comparative CT method requires the target gene and the control gene have approximately equal efficiency, while the relative standard curve method does not require equal efficiency. In the present experiment, the relative standard curve method was used to correct for variation in efficiency. Detailed information on adipocyte RNA isolation and
QPCR can be found in Section 2.5 and Section 2.7 respectively.
Results: QPCR demonstrated that PDE3B and PDE4 (A, B, C, D) genes were detected
in rat adipocyte RNA samples (n=5) and rat heart tissue RNA sample (n=1). In rat
adipocytes, the level of PDE3B, PDE4A, PDE4B, PDE4C, and PDE4D normalized to
28S rat RNA (mean ± SEM) was 38.9 ± 6.6, 2.9 ± 0.9, 7.6 ± 2.4, 7.4 ± 1.8, and 2.5 ± 0.4,
respectively (Figure 4.3). The level of PDE4A, PDE4B, PDE4C, and PDE4D relative to
that of PDE3B was 7%, 18.7%, 18.9%, and 7.2%, respectively. In heart tissue sample, the
level of PDE3B, PDE4A, PDE4B, PDE4C, and PDE4D normalized to 28S rat RNA was
1.4, 4.7, 1.8, 7.9, and 1.7, respectively (Figure 4.4).
4.4 PDE3B and PDE4B protein expression in rat adipocytes
Rationale: It was shown that four PDE4 subtype genes were expressed in rat
adipocytes, so it is expected that all PDE4 subtypes are expressed at the protein level.
Commercial PDE4B and PDE4D antibodies are available, but the specificity of PDE4D
antibody was very poor in preliminary experiments. Because this PDE4B antibody was
generated using a C-terminal peptide common to all PDE4B isoforms, this PDE4B
antibody is expected to detect the expression of all PDE4B variants.
73 Hypothesis: If PDE4B gene is expressed in rat adipocytes, then PDE4B protein will be
present.
Experimental design: Based on the results of preliminary experiments, the specificity
of PDE3B antibody and PDE4B antibody was poor. Therefore, preimmune rabbit serum
was used as a control in this experiment. Proteins in both the particulate and cytosolic
fractions of rat adipocytes were subjected to SDS-PAGE and Western blot. Detailed
information on preparation of adipocyte subcellular fractions and Western blot for
PDE3B and PDE4B expression can be found in Section 2.8.1 and Section 2.8.3 respectively.
Results: Using PDE3B N-terminal antibody, one band with approximate molecular
weight of 135 kDa was identified in the particulate fraction (Figure 4.5). No significant band was detected at the corresponding site of the preimmune control. This 135 kDa band was tentatively identified as PDE3B. Using PDE4B C-terminal antibody, one band with approximate molecular weight of 92 kDa was identified in the cytosolic fraction (Figure
4.6). No significant band was detected at the corresponding site of the preimmune control.
According to molecular weight, this 92 kDa band was tentatively identified as PDE4B3.
4.5 Discussion
The present study characterized PDE4 expression in rat adipocytes. This is the first report to determine and quantitate the expression of PDE4 subtypes in rat adipocytes. The results showed that all four PDE4 subtypes (A, B, C, D) were expressed in rat adipocytes, with PDE4B and PDE4C predominant.
74 PDE4 is expressed by four genes (4A, 4B, 4C, 4D) (Bolger, 1994). By RT-PCR, it
has been known that PDE4A5, PDE4B2, PDE4C2, PDE4D3 and PDE4D5 were
expressed in 3T3-F442A preadipocytes (MacKenzie et al., 1998). In the present study, the
results of both RT-PCR and QPCR demonstrated that all four PDE4 subtypes were
expressed in rat adipocytes. The level of PDE3B was about 14 times higher than that of
PDE4A and PDE4D and was about 5 times higher than that of PDE4B and PDE4C in rat
adipocytes. The average levels of PDE4B and PDE4C were nearly three-fold higher than those of PDE4A and PDE4D in rat adipocytes. The total relative expression of all four
PDE4 subtypes was around 50% of that of PDE3B in rat adipocytes.
In the present study, heart tissue mRNA was used as a positive control. The level of
PDE3B in heart tissue was much lower than in adipocytes. However, the levels of PDE4 subtypes in heart tissue were comparable to that in adipocytes. PDE4C was not detected by RT-PCR in rat heart tissue in the previous study (Kostic et al., 1997). However,
PDE4C was detected in the present study by QPCR. This inconsistency may result from the use of different primers for amplification of PDE4C. In the present study, the efficiency for PDE4C was much lower than another three PDE4 subtypes, suggesting possible difficulty in amplifying PDE4C.
The technique of QPCR has been widely used in research as a sensitive and reliable molecular technique for investigating gene expression. When comparing the expression of different genes using QPCR, equivalent reverse-transcription efficiency is a requirement in the absence of quantitative standards such as purified RNA transcripts. In the present study, the assumption of equivalence was supported because the primer sets were designed to a region of homology across the five genes. The underlying belief was
75 that similar mRNA sequences would have similar secondary structures and thus equivalent reverse-transcription efficiency.
Based on the quantitative data of PDE4 gene expression and the availability of
PDE4 subtype antibodies, PDE4B protein expression was determined in the present study.
Three PDE4B splice variants have been identified in primary rat cortical neurons,
including two long isoforms, PDE4B1 (110 kDa) and PDE4B3 (92 kDa), and one short
isoform, PDE4B2 (65 KDa) (D'Sa et al., 2002). It was reported that all three human
PDE4B isoforms were present in both the cytosolic and the particulate fractions of
transfected COS-7 cells. The distribution proportions of PDE4B1, PDE4B2, and PDE4B3
in cytosol were 71%, 61%, and 58%, respectively (Huston et al., 1997). In the present
study, the PDE4B antibody recognized PDE4B3 in the cytosolic fraction but not in the
particulate fraction of rat adipocytes. In light of the poor specificity of this PDE4B
antibody, it is possible that PDE4B3 may be present in the particulate fraction of rat
adipocytes.
Detection of PDE3B in the particulate fraction in the present study was consistent
with the previous studies with rat adipocytes (Degerman et al., 1998; Liu and Maurice,
1998). Many efforts were made to improve the conditions of Western blot, including
adjusting antibody dilution ratio, extending antibody incubation time, increasing the
protein content of loading samples, changing blocking solution, and purifying protein by immunoprecipitation. However, the results of Western blot were still unsatisfactory because of the poor quality of antibody. Any information other than the PDE4B3 expression in the cytosolic fraction of rat adipocytes can not be interpreted from the results of Western blot. Some bands besides these 135 kDa and 92 kDa bands disappeared
76 in the preimmune control. A possible reason was that the concentration of IgG in
preimmune rabbit serum was lower than that in PDE antibody, so some bands may not
have been recognized. Therefore, it is possible that 135 kDa and 92 kDa bands disappeared in the preimmune control as result of the low concentration of IgG.
The significance of differential expression of PDE4 isoforms in a variety of organs, tissues, and cells remains unclear. Knockout studies provide a possibility for answering this question (Ariga et al., 2004; Mehats et al., 2002). Ablation of PDE4B and PDE4D,
but not PDE4A, significantly reduced neutrophil recruitment in a model of mouse lung
injury induced by endotoxin inhalation. Moreover, the subsequent inhibition of PDE4 by
rolipram had additional inhibitory effects on neutrophil recruitment in PDE4B(-/-) and
PDE4D(-/-) mice respectively, suggesting that the functions of PDE4B and PDE4D were
complementary, but not redundant, in the regulation of neutrophil function (Ariga et al.,
2004). The physiological significance of differential expression of four PDE4 subtypes in
rat adipocytes is unknown.
In summary, four PDE4 subtypes were expressed in rat adipocytes with PDE4B and
PDE4C predominant. The total relative expression of all four PDE4 isoforms was
approximately 50% of that of PDE3B in rat adipocytes. PDE4B3 was detected in the
cytosolic fraction of rat adipocytes. The present study further confirms the lipolysis data
of the ginseng study using the PDE4 inhibitor. If better antibodies for four PDE4
subtypes had been available, the protein expression and subcellular distribution of PDE4
subtypes in rat adipocytes may have been further clarified.
77
PDE3 PDE4
Old nomenclature cGMP-inhibited PDE cAMP specific PDE
Genes A, B A, B, C, D
Substrates cAMP, cGMP cAMP
Km (µM) 0.1-0.8 1-3
Inhibitors cilostamide rolipram OPC3911 Ro 20-1724 milrinone RP73401 amrinone zadarverine
Regulation inhibited by cGMP phosphorylated and activated phosphorylated and activated by PKA by PKB phosphorylated by ERK phosphorylated and activated by PKA
Intracellular particulate fraction particulate fraction Distribution (PDE3B) (PDE4A1) cytosolic fraction cytosolic fraction (PDE3A) (PDE4D1, 4D2) particulate and cytosolic fraction (PDE4A4, 4A5, 4A8; PDE4B1, 4B2, 4B3; PDE4D3, 4D4, 4D5)
Organ and Tissue heart, brain, heart, liver, lung, Distribution vascular smooth muscle, kidney, immune cells, platelets, adipose tissue, blood cells, preadipocytes liver, pancreas
Table 4.1. Comparison of PDE3 and PDE4.
78 Gene Amplicon (Genbank Accession Sequence (bp) No.)
PDE3B F, CAGGAAGGATTCTCAGTCAG 530 (NM_017229) R, GTATTCTGGGCGAGAAAGAT
PDE4A F, GCGGGACCTACTGAAGAAATTCC 233 (NM_013101) R, CAGGGTGGTCCACATCGTGG
PDE4B F, CAGCTCATGACCCAGATAAGTGG 786 (NM_017031) R, GTCTGCACAATGTACCATGTTGCG
PDE4C F, ACTGAGTCTGCGCAGGATGG 539 (XM_214325) R, CCTCCTCTTCCTCTGTCTCCTC
PDE4D F, CCTCTGACTGTTATCATGCACACC 262 (NM_017032) R, GATCCACATCATGTATTGCACTGGC
Table 4.2. Primer pairs for RT-PCR to detect sequences encoding PDE3B and PDE4 (A, B, C, D). PDE3B and PDE4 (A, B, C, D) primers were described by Harndahl (Harndahl et al., 2002) and Kostic (Kostic et al., 1997), respectively. F, forward primer; R, reverse primer.
79
RT (+) ( RT RT (+) ( RT RT (+) ( RT RT (+) ( RT RT (+) ( RT
− − − − − ) ) ) ) )
80 PDE3B PDE4A PDE4B PDE4C PDE4D (530 bp) (233 bp) (786 bp) (539 bp) (262 bp)
Figure 4.1. Expression of PDE3B and PDE4 (A, B, C, D) in rat adipocytes. Total RNA extracted from isolated rat adipocytes was reverse-transcribed, and cDNA was amplified using specific primers for PDE3B and PDE4 (A, B, C, D). The predicted fragments for PDE3B (530 bp), PDE4A (233 bp), PDE4B (786 bp), PDE4C (539 bp), and PDE4D (262 bp) were amplified by PCR. Experiments were done in the presence (+) or absence (–) of reverse transcriptase (RT). Results shown are representative of three independent experiments. The first lane in each panel is DNA ladder.
80
Query: 1 actttatttcaagatactgggtttattgggaaacatttaaaattcccactcaagaattta 60 |||||||||||||||||||| |||||||| |||||||||||||||||||||||||||||| Sbjct: 2114 actttatttcaagatactgg-tttattgg-aaacatttaaaattcccactcaagaattta 2171
Query: 61 tgaattattttcgtgcactagaaaatggctaccgagatattccatatcacaatcgtgtgc 120 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2172 tgaattattttcgtgcactagaaaatggctaccgagatattccatatcacaatcgtgtgc 2231
Query: 121 atgccacagatgtcctacatgctgtttggtatttgacaacacgaccaattcctggcttac 180 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2232 atgccacagatgtcctacatgctgtttggtatttgacaacacgaccaattcctggcttac 2291
Query: 181 agcagctccataataaccatgaaacagaaaccaaagcagattcagatgctagacttagtt 240 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2292 agcagctccataataaccatgaaacagaaaccaaagcagattcagatgctagacttagtt 2351
Query: 241 ctggacagattgcttacctttcttcgaagagttgctgtattccagataagagttatggct 300 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2352 ctggacagattgcttacctttcttcgaagagttgctgtattccagataagagttatggct 2411
Query: 301 gcctgtcttcaaacattcctgcgttagaactgatggctttatatgtggcagctgccatgc 360 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2412 gcctgtcttcaaacattcctgcgttagaactgatggctttatatgtggcagctgccatgc 2471
Query: 361 acgattatgatcacccaggaagaacaaatgcattcctagtgggctacaaatgcacctcag 420 |||||||||||||||||||||||||||||||||||||||| ||||||||||||||||||| Sbjct: 2472 acgattatgatcacccaggaagaacaaatgcattcctagt-ggctacaaatgcacctcag 2530
Query: 421 gcagttttatacaatgacagatctgttctagaaaatcatcatgccgcatcagcgtggaat 480 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2531 gcagttttatacaatgacagatctgttctagaaaatcatcatgccgcatcagcgtggaat 2590
Query: 481 ctgtatctttctcgcccnaaanacaa 506 ||||||||||||||||| || |||| Sbjct: 2591 ctgtatctttctcgcccagaatacaa 2616
Table 4.3. Sequence of PDE3B PCR product aligned with the published sequence of PDE3B using NCBI two-sequence BLAST. The sequence denoted by “Query” is PDE3B PCR product. The sequence denoted by “Sbjct” is the published sequence of PDE3B.
81
Query: 3 tgtggacaccatgatgatgtacattgctgaccctggaggaccactaccatgccgacgtgg 62 |||||||||||||||||||||||| ||||||||||||||||||||||||||||||||||| Sbjct: 1280 tgtggacaccatgatgatgtacat-gctgaccctggaggaccactaccatgccgacgtgg 1338
Query: 63 cctaccacaacagcctgcacgcagcggatgtgctgcagtccacacacgtgctgctggcca 122 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1339 cctaccacaacagcctgcacgcagcggatgtgctgcagtccacacacgtgctgctggcca 1398
Query: 123 cgcccgcactggacgctgtgttcacagacctggagattcttgctgccctcttcgctgctg 182 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1399 cgcccgcactggacgctgtgttcacagacctggagattcttgctgccctcttcgctgctg 1458
Query: 183 ccatccacgatgtggaccaccct 205 ||||||||||||||||||||||| Sbjct: 1459 ccatccacgatgtggaccaccct 1481
Table 4.4. Sequence of PDE4A PCR product aligned with the published sequence of PDE4A using NCBI two-sequence BLAST. The sequence denoted by “Query” is PDE4A PCR product. The sequence denoted by “Sbjct” is the published sequence of PDE4A.
82
Query: 9 acagctcaagccttgaacaacacaangcatctcacgctttggagtcaacacggaaaatga 68 ||||||||||||| ||||||||||| |||||||||||||||||||||||||||||||||| Sbjct: 1413 acagctcaagcct-gaacaacacaa-gcatctcacgctttggagtcaacacggaaaatga 1470
Query: 69 ggatcatctagccaaggagctggaagacctgaacaaatggggccttaacatcttcaacgt 128 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1471 ggatcatctagccaaggagctggaagacctgaacaaatggggccttaacatcttcaacgt 1530
Query: 129 ggctgggtactcccataatcggcccctcacatgcatcatgtacgccattttccaggaaag 188 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1531 ggctgggtactcccataatcggcccctcacatgcatcatgtacgccattttccaggaaag 1590
Query: 189 agaccttctaaagacgtttaaaatctcctccgacaccttcgtaacctacatgatgacttt 248 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1591 agaccttctaaagacgtttaaaatctcctccgacaccttcgtaacctacatgatgacttt 1650
Query: 249 agaagaccattaccattctgatgtggcgtatcacaacagcctgcacgctgctgacgtggc 308 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1651 agaagaccattaccattctgatgtggcgtatcacaacagcctgcacgctgctgacgtggc 1710
Query: 309 ccagtcaacgcacgttctcctctctacgccagcactggatgctgtcttcacagacctgga 368 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1711 ccagtcaacgcacgttctcctctctacgccagcactggatgctgtcttcacagacctgga 1770
Query: 369 aatcctggctgccatttttgcagctgccatccatgatgttgatcatcctggagtctccaa 428 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1771 aatcctggctgccatttttgcagctgccatccatgatgttgatcatcctggagtctccaa 1830
Query: 429 tcagtttctcatcaatacaaattccgaacttgctttgatgtataatgacgaatctgtgct 488 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1831 tcagtttctcatcaatacaaattccgaacttgctttgatgtataatgacgaatctgtgct 1890
Query: 489 ggaaaaccatcacctcgctgtgggattcaagctccttcaagaggaacattgcgacatctt 548 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1891 ggaaaaccatcacctcgctgtgggattcaagctccttcaagaggaacattgcgacatctt 1950
Query: 549 tcagaatcttaccaagaagcaacgccagacactcaggaaaatggtgattgacatggtgtt 608 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1951 tcagaatcttaccaagaagcaacgccagacactcaggaaaatggtgattgacatggtgtt 2010
Query: 609 agcaactgatatgtccaagcacatgagcctcctggctga-cttaaaacgatggtagaaac 667 ||||||||||||||||||||||||||||||||||||||| |||||||||||||||||||| Sbjct: 2011 agcaactgatatgtccaagcacatgagcctcctggctgaccttaaaacgatggtagaaac 2070
Query: 668 caaaaaggtgacgagctccggtgttctcctcctggacaactatactgaccggatacaggt 727 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2071 caaaaaggtgacgagctccggtgttctcctcctggacaactatactgaccggatacaggt 2130
Query: 728 tcttcgcaacatggtaca 745 |||||||||||||||||| Sbjct: 2131 tcttcgcaacatggtaca 2148
Table 4.5. Sequence of PDE4B PCR product aligned with the published sequence of PDE4B using NCBI two-sequence BLAST. The sequence denoted by “Query” is PDE4B PCR product. The sequence denoted by “Sbjct” is the published sequence of PDE4B. 83
Query: 13 cacagacattgtccaagcacatggagcctcctaggctgacctcaagaccattggttggag 72 ||||||||| ||||||||||||| |||||||| |||||||||||||||||| | ||||| Sbjct: 1782 cacagacat-gtccaagcacatg-agcctcct-ggctgacctcaagaccatgg--tggag 1836
Query: 73 accaagaaagtgactatgccttggcgtcctgctcttggacaactactctgaccgcatcca 132 |||||||||||||||| ||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1837 accaagaaagtgacta-gccttggcgtcctgctcttggacaactactctgaccgcatcca 1895
Query: 133 ggtcctccagagcctggtgcactgcgccgacctcagcaaccctgccaagccactacccct 192 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1896 ggtcctccagagcctggtgcactgcgccgacctcagcaaccctgccaagccactacccct 1955
Query: 193 ctaccgccagtggacggagcgcatcatggctgagttcttccagcagggtgaccgggaacg 252 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1956 ctaccgccagtggacggagcgcatcatggctgagttcttccagcagggtgaccgggaacg 2015
Query: 253 tgagtcgggcttggacatcagccccatgtgcgacaagcacacagcctcggtggagaaatc 312 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2016 tgagtcgggcttggacatcagccccatgtgcgacaagcacacagcctcggtggagaaatc 2075
Query: 313 ccaggtgggattcattgactacatcgctcacccattgtgggaaacttgggccgacctggt 372 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2076 ccaggtgggattcattgactacatcgctcacccattgtgggaaacttgggccgacctggt 2135
Query: 373 gcaccccgatgcccaggagctgctggataccttggaagacaacagagagtggtatcagag 432 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2136 gcaccccgatgcccaggagctgctggataccttggaagacaacagagagtggtatcagag 2195
Query: 433 tagggttccctgcagccctccacacgccattggccctgacaggttcaagtttgagctgac 492 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 2196 tagggttccctgcagccctccacacgccattggccctgacaggttcaagtttgagctgac 2255
Query: 493 cctggaggagacagaggaanaggagga 519 ||||||||||||||||||| ||||||| Sbjct: 2256 cctggaggagacagaggaagaggagga 2282
Table 4.6. Sequence of PDE4C PCR product aligned with the published sequence of PDE4C using NCBI two-sequence BLAST. The sequence denoted by “Query” is PDE4C PCR product. The sequence denoted by “Sbjct” is the published sequence of PDE4C.
84
Query: 1 tttcaggaacgagatttgttaaaaacgtttaaaatcccagtggacactttgattacgtat 60 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 874 tttcaggaacgagatttgttaaaaacgtttaaaatcccagtggacactttgattacgtat 933
Query: 61 cttatgactctagaagaccattaccatgctgacgtggcctatcacaacaacatccatgct 120 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 934 cttatgactctagaagaccattaccatgctgacgtggcctatcacaacaacatccatgct 993
Query: 121 gcagatgtcgtccagtcaactcatgtgctgctctctacacccgctttggaggctgttttc 180 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 994 gcagatgtcgtccagtcaactcatgtgctgctctctacacccgctttggaggctgttttc 1053
Query: 181 actgacttggagattctcgcggccatttttgccagtgcaatacatgatgtggatca 236 |||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct: 1054 actgacttggagattctcgcggccatttttgccagtgcaatacatgatgtggatca 1109
Table 4.7. Sequence of PDE4D PCR product aligned with the published sequence of PDE4D using NCBI two-sequence BLAST. The sequence denoted by “Query” is PDE4D PCR product. The sequence denoted by “Sbjct” is the published sequence of PDE4D.
85
1 90 GTGACNAGCT CTGGCGTTCT CCTCCTGGAC AACTANTCTG ACCGNATCCA GGTCCTCCAG AACATGGTGC ACTGTGCAGA CCTCAGCAAC 4A ...... GGAC AACTACTCTG ACCGTATCCA GGTCCTCAGG AACATGGTGC ACTGTGCAGA CCTCAGCAAT 4B GTGACGAGCT CCGGTGTTCT CCTCCTGGAC AACTATACTG ACCGGATACA GGTTCTTCGC AACATGGTAC ATTGTGCAGA CCTGAGCAAC 4C ...... 4D ...... CCTCCAG AATATGGTGC ACTGTGCAGA CCTGAGCAAC 3B ...... AGC AGTGAAAACG ATCGACTCTT AGTCTGCCAG GTGTGCATCA AATTAGCAGA CATCAACGGC
91 150 190 CCNACCAAGC CNCTGGAGCT NTACCGNCAG TGGACNGANC GCATCATGGA TGAGTTCTTC CAGCAGGGNG ACCGAGAACG NGAGCGGGGN ATGGAGATN CCCACCAAGC CCCTGGAGCT GTACCGACAG TGGACCGAC...... CCTACCAAGT CCTTGGAGTT GTATCGGC...... GGACGGAGC GCATCATGGC TGAGTTCTTC CAGCAGGGTG ACCGGGAACG TGAGTCGGGC TTGGACATC. CCCACAAAGC CACTCCAGCT CTACCGCCAG TGGACGGACC GGATAATGGA GGAGTTCTTC CGTCAGGGGG ACCGGGAGCG TGAGCGTGGC ATGGAGATA. CCAGCAAAAG ATCGGGATCT TCATTTGAGA TGGACAGAAG GCA......
86
Table 4.8. Alignment of PDE3B and PDE4 (A, B, C, D) primers and probes for QPCR. Red, forward primer; green, probe; blue, reverse primer.
86
NM_017229 PDE3B(2736)-----CCAATGATGTAAATAG---TAACGGTATAGAATGGAGCAGTGAAA XM_214325 PDE4C(1833) GGAGACCAAGAAAGTGACTAGCCTTGGCGTCCTGCTCTTGGACAACTACT NM_013101 PDE4A(1727) GGAGACGAAGAAAGTGACCAGCTCCGGAGTTCTCTTGCTGGACAACTACT NM_017031 PDE4B(2065) AGAAACCAAAAAGGTGACGAGCTCCGGTGTTCTCCTCCTGGACAACTATA NM_017032 PDE4D(1359) TGAAACGAAGAAGGTGACGAGCTCTGGCGTCCTCCTCCTTGATAACTATT Consensus GA ACCAAGAA GTGAC AGCTCTGGCGTTCTCCTCCTGGACAACTA T
NM_017229 PDE3B(2778) ACGATCGACTCTTAGTCTGCCAGGTGTGCATCAAATTAGCAGACATCAAC XM_214325 PDE4C(1883) CTGACCGCATCCAGGTCCTCCAGAGCCTGGTGCACTGCGCCGACCTCAGC NM_013101 PDE4A(1777) CTGACCGTATCCAGGTCCTCAGGAACATGGTGCACTGTGCAGACCTCAGC NM_017031 PDE4B(2115) CTGACCGGATACAGGTTCTTCGCAACATGGTACATTGTGCAGACCTGAGC NM_017032 PDE4D(1409) CTGACAGGATCCAGGTCCTCCAGAATATGGTGCACTGTGCAGACCTGAGC Consensus CTGACCG ATCCAGGTCCTCCAGAACATGGTGCACTGTGCAGACCTCAGC
NM_017229 PDE3B(2828) GGCCCAGCAAAAGATCGGGATCTTCATTTGAGATGGACAGAAGGCATTGT XM_214325 PDE4C(1933) AACCCTGCCAAGCCACTACCCCTCTACCGCCAGTGGACGGAGCGCATCAT NM_013101 PDE4A(1827) AATCCCACCAAGCCCCTGGAGCTGTACCGACAGTGGACCGACCGCATCAT NM_017031 PDE4B(2165) AACCCTACCAAGTCCTTGGAGTTGTATCGGCAATGGACTGATCGCATCAT NM_017032 PDE4D(1459) AACCCCACAAAGCCACTCCAGCTCTACCGCCAGTGGACGGACCGGATAAT Consensus AACCC ACCAAGCC CTGGAGCT TACCG CAGTGGAC GA CGCATCAT
Table 4.9. A region of high similarity among PDE3B and PDE4 (A, B, C, D) sequences.
87 32 y = -3.42x + 30.61 y = -4.52x + 33.09 y = -3.47x + 32.31 27 y = -3.55x + 30.21
y = -3.29x + 28.81 22
PDE4A PDE4B PDE3B PDE4D PDE4C 28s Ct (4A 4B) 28s Ct (3B) 28s Ct (4C 4D) Linear (PDE4B) Linear (PDE4A) Linear (PDE4C) Linear (PDE4D) Linear (PDE3B) Linear (28s Ct (4A 4B)) Linear (28s Ct (3B)) 17 Linear (28s Ct (4C 4D)) Threshold Cycle (Ct) Cycle Threshold y = -4.4x + 18.41 y = -3.93x + 17.49 12
y = -3.94x + 17.30
7 0.0 1.0 2.0
Log10 ng Total RNA
Figure 4.2. The standard curves for the amplification of PDE3B and PDE4 (A, B, C, D). Three calibration curves were generated using serially diluted samples of 28s rat RNA for each target gene. The equation is shown for each standard curve. Total RNA samples were from 5 independent adipocyte preparations. Each QPCR sample was amplified in triplicate. Threshold cycle (Ct) is the cycle number at which the fluorescence generated within a reaction crosses the threshold.
88 Standard curve Gene Amplicon equation (Genbank Sequence Efficiency (bp) (correlation Accession No.) coefficient)
PDE3B 2768F, AGCAGTGAAAACGATCGACTCTT (NM_017229) 2795T, TGCCAGGTGTGCATCAAATTAGCAGAC 106 y= -3.47x + 32.31 94.18% 2873R, TGCCTTCTGTCCATCTCAAATG (0.99)
PDE4A 1766F, GGACAACTACTCTGACCGTATCCA (NM_013101) 1821T, GTCGGTCCACTGTCGGTACA 103 y= -3.29x + 28.81 101.18% 1868R, CTCAGCAATCCCACCAAGCCCCT (0.995)
PDE4B 2078F, GTGACGAGCTCCGGTGTTC (NM_017031) 2100T, TCCTGGACAACTATACTGACCGGATACAGGTT 118 y= -3.42x +30.61 96.21% 2195R, GCCGATACAACTCCAAGGACTT (0.992)
PDE4C 1967F, GGACGGAGCGCATCATG (XM_214325) 1985T, CTGAGTTCTTCCAGCAGGGTGACCG 68 y= -4.52x + 33.10 66.42% 89 2034R, GATGTCCAAGCCCGACTCA (0.974)
PDE4D 1425F, CCTCCAGAATATGGTGCACTGT (NM_017032) 1489T, CAGTGGACGGACCGGATAATGGAGG 135 y= -3.55x + 30.21 91.46% 1560R, TATCTCCATGCCACGCTCA (0.996)
Table 4.10. Properties of QPCR amplicons used for relative quantitation of PDE4 (A, B, C, D) gene expression. Primers and probes were designed using the Primer Express software program (Applied Biosystems; Foster City, CA) to a region of high similarity among PDE3B and PDE4 (A, B, C, D) sequences (Table 4.9). F, forward primer; R, reverse primer; T, TaqMan probe.
89
50
40
30
20 Relative Amount Relative 10
0 PDE3B PDE4A PDE4B PDE4C PDE4D
Figure 4.3. Relative expression of PDE3B and PDE4 (A, B, C, D) normalized to 28s RNA in rat adipocytes (n=5). The level of PDE3B was about 14 times higher than that of PDE4A and PDE4D and was about 5 times higher than that of PDE4B and PDE4C. Relative expression levels were determined by the Relative Standard Curve Method (Applied Biosystems; Foster City, CA).
90
10
8
6
4 Relative Amount Relative 2
0 PDE3B PDE4A PDE4B PDE4C PDE4D
Figure 4.4. Relative expression of PDE3B and PDE4 (A, B, C, D) normalized to 28s RNA in rat heart tissue (n=1). The level of PDE3B was comparable to PDE4B and PDE4D, with PDE4A and PDE4C predominant. Relative expression levels were determined by the Relative Standard Curve Method (Applied Biosystems; Foster City, CA).
91
A B
132 kDa 132 kDa
78 kDa 78 kDa
C P C P
Figure 4.5. Expression of PDE3B in rat adipocytes. Cytosol fraction (C) and particulate fraction (P) were prepared as described in Section 2.8.1. Protein (20 µg) was subjected to SDS-PAGE and Western blot was performed using PDE3B N-terminal antibody (A) and preimmune rabbit serum (B). Results shown were representative of 3 independent experiments. The band with approximate molecular weight of 135 kDa (denoted with a block arrow) was identified by PDE3B N-terminal antibody in the particulate fraction of adipocytes. There was no significant band at the corresponding site of the preimmune control.
92
A B
97 kDa 97 kDa 66 kDa 66 kDa
C P C P
Figure 4.6. Expression of PDE4B3 in rat adipocytes. Cytosol fraction (C) and particulate fraction (P) were prepared as described in Section 2.8.1. Protein (20 µg) was subjected to SDS-PAGE and Western blot was performed using PDE4B C-terminal antibody (A) and preimmune rabbit serum (B). Results shown were representative of 3 independent experiments. The band with approximate molecular weight of 92 kDa (denoted with a block arrow) was identified by PDE4B C-terminal antibody in the cytosol fraction of adipocytes. There was no significant band at the corresponding site of the preimmune control.
93
CHAPTER 5
THE ROLE OF PHOSPHODIESTERASE 4 IN ANTILIPOLYSIS MEDIATED
BY PROSTAGLADIN E2 AND ADENOSINE IN RAT ADIPOCYTES IN VITRO
5.1 Introduction
The ginseng study showed that ginseng antilipolysis is mediated in part by PDE4.
Also, the signaling pathway for ginseng antilipolysis was shown to be distinct from that of insulin antilipolysis, although the specific pathway for the former remains unknown. It is of interest to investigate how PDE4 is activated in adipocytes. The multiple components make it difficult to investigate the signaling pathway for ginseng antilipolysis. If an endogenous factor has a similar pattern of antilipolytic effect as ginseng, investigation of the signaling pathway used by the endogenous antilipolytic factor may provide insights into the mechanism responsible for the antilipolytic effect of ginseng.
Besides insulin, PGE2 and adenosine are two potent endogenous antilipolytic agents released by adipocytes (Vassaux et al., 1992). An early study compared the mechanisms by which insulin, PGE2 and adenosine inhibited basal lipolysis and lipolysis stimulated by multiple agents in isolated human adipocytes (Lonnqvist et al.,
1989). The following lipolytic agents stimulated lipolysis at different steps: isoproterenol (a β-adrenergic receptor agonist); cholera toxin (a Gs protein activator); pertussis toxin (a Gi protein inhibitor); forskolin (an adenylate cyclase activator); 94 enprofylline (a nonselective PDE inhibitor); N6-monobutyryl cAMP (a nonhydrolyzable cAMP analog); 8-bromo cAMP (a long-acting derivative of cAMP analog). PGE2 and PIA inhibited basal lipolysis and lipolysis stimulated by isoproterenol, forskolin, enprofylline, N6-monobutyryl cAMP, and 8-bromo cAMP, but they did not inhibit cholera toxin- or pertussis toxin-stimulated lipolysis. In contrast, insulin inhibited basal lipolysis and lipolysis stimulated by isoproterenol, forskolin, cholera toxin, pertussis toxin and 8-bromo cAMP. However, insulin did not inhibit lipolysis stimulated by enprofylline and N6-monobutyryl cyclic AMP. The results suggested that insulin and adenylate cyclase inhibitors inhibited lipolysis by different mechanisms. Insulin inhibited lipolysis by activating PDE, while PGE2 and
PIA inhibitors inhibited lipolysis by regulating G proteins. The above study also suggested that PGE2 and PIA might inhibit lipolysis by other mechanisms. Because
PGE2 and PIA inhibited lipolysis stimulated by an adenylate cyclase activator, a PDE inhibitor, and cAMP analogs, it is possible that PGE2 and PIA inhibit lipolysis through activating PDE or suppressing hormone-sensitive lipase.
Since the previous study proposed the possible role of PDE in the antilipolytic effects of PGE2 and PIA, the present study further determined if PDE mediates PGE2 and adenosine antilipolysis in rat adipocytes.
5.2 Effects of insulin and PGE2 on lipolysis
Rationale: Endogenous antilipolysis can be mediated by insulin, PGE2, and adenosine. It was reported that PGE1 increased PDE activity in human adipose tissue
(Engfeldt et al., 1983). Therefore, PGE2 may also increase PDE activity in rat adipocytes and thus inhibit lipolysis.
95 Hypothesis: If PGE2 inhibits lipolysis by activating PDE in part, then specific
PDE3 or PDE4 inhibitors will reduce PGE2 antilipolysis partially.
Experimental design: Freshly isolated adipocytes were incubated with the specific
PDE3 inhibitor cilostamide (5 µM), the specific PDE4 inhibitor rolipram (10 µM), and the combination of cilostamide (5 µM) and rolipram (10 µM) in the absence and presence of PGE2 (10 nM). The concentration of PGE2 was based on a previous study
(Lonnqvist et al., 1989). Detailed information on rat adipocyte isolation and lipolysis assay can be found in Section 2.3 and Section 2.4 respectively.
Results: Insulin inhibited lipolysis by 45.1% compared to basal (n=5; P<0.006;
Figure 5.1). The PDE3 inhibitor, cilostamide, completely reversed insulin antilipolysis compared to basal with cilostamide. The PDE4 inhibitor, rolipram, did not reduce insulin antilipolysis.
PGE2 inhibited lipolysis by 84% compared to basal (n=5; P<0.006; Figure 5.1).
Cilostamide reduced PGE2 antilipolysis to 76.3% compared to basal with cilostamide, and PGE2 inhibited lipolysis (P<0.006). Rolipram reduced PGE2 antilipolysis to
46.6% compared to basal with rolipram, and PGE2 still inhibited lipolysis (P<0.006).
The combination of two PDE inhibitors reduced PGE2 antilipolysis to 17.5% compared to basal with two inhibitors, with a synergistic effect defined as greater than the expected addition of each inhibitor alone (P<0.02).
5.3 Effects of insulin and PIA on lipolysis
Rationale: PDE4 contributed to the antilipolytic effects of PGE2 and ginseng. Both
PGE2 and adenosine can inhibit adenylate cyclase. Therefore, PDE4 may also mediate
PIA antilipolysis.
96 Hypothesis: If PIA inhibits lipolysis by activating PDE in part, then specific PDE3 or PDE4 inhibitors will partially reduce PIA antilipolysis.
Experimental design: Freshly isolated adipocytes were incubated with the specific
PDE3 inhibitor cilostamide (5 µM), the specific PDE4 inhibitor rolipram (10 µM), and combination of cilostamide (5 µM) and rolipram (10 µM) in the absence and presence of PIA (a nonhydrolyzable analog of adenosine, 10 nM). The concentration of PIA was based on a previous study (Smith and Manganiello, 1989). Detailed information on rat adipocyte isolation and lipolysis assay can be found in Section 2.3 and Section 2.4 respectively.
Results: Insulin inhibited lipolysis by 28.1% compared to basal (n=5; P<0.006;
Figure 5.2). As seen before, the PDE3 inhibitor, cilostamide, completely reversed insulin antilipolysis compared to basal with cilostamide. The PDE4 inhibitor rolipram did not reduce insulin antilipolysis.
PIA inhibited lipolysis by 92.2% compared to basal (P<0.006). Neither cilostamide nor rolipram alone reduced PIA antilipolysis. The combination of the two inhibitors reduced PIA antilipolysis to 56.8% of basal with the two inhibitors; this effect was synergistic (P<0.02).
5.4 Effects of PDE inhibitors on basal lipolysis
Rationale: cAMP is a key molecule in the regulation of lipolysis. Both PDE3 and
PDE4 hydrolyze cAMP to AMP, and are present in rat adipocytes. Specific inhibition of PDE3 and PDE4 increased the rate of lipolysis in 3T3-L1 adipocytes and cultured rat adipocytes (Elks and Manganiello, 1984; Snyder et al., 2005). Specific inhibitors of PDE3 and PDE4 increased basal lipolysis in the ginseng, PGE2, and PIA studies.
97 Hypothesis: If basal lipolysis is increased by specific inhibitors of PDE3 and PDE4, then combined inhibition of PDE3 and PDE4 will increase basal lipolysis in an additive manner.
Experimental design: 18 independent experiments on basal lipolysis were summarized from the ginseng, PGE2, and PIA studies. In the present study, the basal condition is defined as an adenosine-free system. Given that different adipocyte preparations may have varied rates of lipolysis due to differences in release of endogenous adenosine from disrupted adipocytes (Honnor et al., 1985). Adenosine deaminase (0.8 U/ml) was added to adipocyte incubations to minimize variations in basal lipolysis and sufficiently maintain basal lipolysis to measure the antilipolytic response to insulin, PGE2, and PIA.
Results: The PDE3 inhibitor cilostamide and the PDE4 inhibitor rolipram increased basal lipolysis by 28% and 33%, respectively (n=18, P<0.001, Figure 5.3). The combination of cilostamide and rolipram increased basal lipolysis by 51%, higher than cilostamide or rolipram alone (P<0.001), but the effect were slightly less than additive.
5.5 Discussion
The present study examined the role of PDE3B and PDE4 in antilipolysis mediated by the endogenous inhibitors of adenylate cyclase, PGE2 and PIA. The interesting result was that the PDE4 inhibitor rolipram alone reduced PGE2 antilipolysis, but not PIA antilipolysis. Moreover, combination of the PDE3 and
PDE4 inhibitors reduced PGE2 and PIA antilipolysis in a synergistic manner.
Previous studies showed that inhibition of PDE4 increased the rate of lipolysis in
3T3-L1 adipocytes (Elks and Manganiello, 1984; Snyder et al., 2005) and rat 98 adipocytes (Schmitz-Peiffer et al., 1992; Shechter, 1984; Snyder et al., 2005). In contrast with PDE3B, insulin antilipolysis was not reversed by the PDE4 inhibitor rolipram. Thus, activation of PDE4 is not thought to play a role in the antilipolytic effect of insulin (Hagstrom-Toft et al., 1995; Schmitz-Peiffer et al., 1992; Shechter,
1984). In agreement with these previous studies, the present study showed that cilostamide, but not rolipram, completely reversed insulin antilipolysis. Of particular interest, rolipram alone reduced PGE2 antilipolysis, suggesting that PDE4 was activated in PGE2 antilipolysis. However, PIA antilipolysis was not reduced by rolipram alone. It is likely that PGE2 antilipolysis is mediated by multiple intracellular mechanisms. Besides inhibiting adenylate cyclase, PGE2 may inhibit lipolysis in rat adipocytes by activation of PDE4 in part.
The summary of three antilipolysis experiments indicated that the PDE3 inhibitor cilostamide and the PDE4 inhibitor rolipram increased basal lipolysis by
28% and 33%, respectively. Based on the previous and present study, it is reasonable to conclude that PDE4 plays a tonic role in antilipolysis along with PDE3B in rat adipocytes.
Synergistic effects of PDE3 and PDE4 inhibitors on lipolysis in cultured rat adipocytes have been reported (Snyder et al., 2005). In consistent with Snyder et al.’s study, the present study showed there was a partially additive effect on basal lipolysis in rat adipocytes. This difference may result from the different response of freshly isolated rat adipocytes in the present study and cultured rat adipocytes in their study.
The shorter incubation time used in the present study (1 hr vs 6 hr) may also contribute to this difference. Moreover, the addition of adenosine deaminase to the incubation medium in the present study represents another noteworthy difference.
99 In the present study, the PDE3 and PDE4 inhibitors synergistically reduced antilipolysis mediated by PGE2 and PIA. It has been reported that the combination of
PDE3 and PDE4 inhibitors has a synergistic effects on cell-specific physiological functions, such as inhibiting vascular smooth muscle cell migration (Palmer et al.,
1998) and decreasing T-lymphocyte proliferation (Bielekova et al., 2000). The proposed mechanism is that this synergism results from the increase in cAMP by inhibiting both PDE3 and PDE4. The synergistic effect on lipolysis in the present study may also be related to the intracellular concentration of cAMP. In the basal state, the intracellular concentration of cAMP was presumed to be high because of addition of adenosine deaminase. After adding PGE2 or PIA, the intracellular concentration of cAMP is expected to decrease due to reduced synthesis of cAMP by inhibition of adenylate cyclase. When PDE3 and PDE4 activities are both blocked, the concentration of cAMP and the lipolysis rate increase.
In summary, the present results demonstrated that both PGE2 and PIA antilipolysis were mediated by PDE. However, the mechanisms by which PGE2 and
PIA act on PDE respectively differed. PGE2 may inhibit lipolysis by activating PDE4 in part. Thus, the mechanisms for PGE2 and ginseng antilipolysis are similar.
100
CON CIL ROL CIL+ROL
500 M) µ 400 * 300
200 * * * 100 * *
Glycerol Concentration ( Concentration Glycerol 0 Basal Insulin PGE2
Figure 5.1. Effects of insulin and PGE2 on lipolysis in rat adipocytes. Rat adipocytes were incubated without (basal) and with insulin (90 pM) and PGE2 (10 nM) in the absence and presence of the PDE3 inhibitor cilostamide (CIL, 5 µM), the PDE4 inhibitor rolipram (ROL, 10µM), and combination of these two inhibitors. Basal indicates that no insulin or PGE2 was present. Control (CON) indicates that no PDE inhibitor was present. Data are mean ± SEM of 5 independent experiments assayed in triplicate. Statistical significance compared to basal for each inhibitor is denoted with *, P <0.006.
101
CON CIL ROL CIL+ROL
500 M) µ 400
300 * * 200 *
100 * * *
Glycerol Concentration ( Concentration Glycerol 0 Basal Insulin PIA
Figure 5.2. Effects of insulin and PIA on lipolysis in rat adipocytes. Rat adipocytes were incubated without (basal) and with insulin (90 pM) or PIA (10 nM) in the absence and presence of the PDE3 inhibitor cilostamide (CIL, 5 µM), the PDE4 inhibitor rolipram (ROL, 10µM), and the combination of these two inhibitors. Basal indicates that no insulin or PIA was present. Control (CON) indicates that no PDE inhibitor was present. Data are mean ± SEM of 5 independent experiments assayed in triplicate. Statistical significance compared to basal for each inhibitor is denoted with *, P <0.006.
102
500 M)
µ * 400 * * 300
200
100 Glycerol Concentration ( 0 CON CIL ROL CIL+ROL
Figure 5.3. Effects of the PDE3 and PDE4 inhibitors on basal lipolysis in rat adipocytes. Rat adipocytes were incubated with the PDE3 inhibitor cilostamide (CIL, 5 µM), the PDE4 inhibitor rolipram (ROL, 10µM), and the combination of these two inhibitors. Control (CON) indicates that no PDE inhibitor was present. Data are presented as mean ± SEM of 18 independent experiments assayed in triplicate. Statistical significance compared to control is denoted with *, P <0.05.
103
CHAPTER 6
ACTIVATION OF PHOSPHODIESTERASE 4 IN RAT ADIPOCYTES
IN VITRO
6.1 Introduction
Lipolysis in adipocytes is regulated by the cAMP system. Because the degradation of cAMP is dependent on PDE, activation of PDE is a key mechanism in the regulation of lipolysis. It has been shown that many lipolytic and antilipolytic agents alter PDE activity in human adipose tissue (Engfeldt et al., 1983). After 100 mg of human adipose tissue fragments were incubated with various agents for 10 min, the results of PDE activity assay showed that insulin (500 µU/ml), isoproterenol (ISO,
6 µM), noradrenaline (5 µM), prostaglandin E1 (PGE1, 0.56mM), and cAMP (10 µM) increased the total PDE activity by 30%, 23%, 27%, 34%, and 41%, respectively.
Adenosine (10 µM), nicotinic acid (1, 100, and 1000 µM), and phentolamine (10 and
100 µM) did not change the total PDE activity. Theophylline (1 mM) decreased the total PDE activity by 20%.
In addition to insulin, ISO also activates PDE3B to degrade cAMP while it stimulates lipolysis. This feedback regulation is helpful for the maintenance of cAMP steady state and control of lipolysis (Shakur et al., 2001). An early study showed that
ISO (100 nM) increased the particulate PDE3B activity by 100% and PKA activity by
104 8 to 10-fold in rat adipocytes after 15 min incubation. Insulin (0.1-0.3 nM) increased the particulate PDE3B activity by 50% without changing PKA activity after 12 to 16 min incubation. In the presence of ISO and insulin, there was a short synergistic effect on stimulation of the particulate PDE3B (Smith and Manganiello, 1989). This study suggests that ISO and insulin activate PDE3B by different mechanisms, and the synergistic effect of ISO and insulin may play a key role in the antilipolytic effect of insulin. Subsequent studies demonstrated that ISO phosphorylates PDE3B at Ser-302 by PKA and insulin phosphorylates PDE3B at Ser-302 by PKB (Rahn et al., 1996;
Smith et al., 1991). This dual phosphorylation of PDE3B may contribute to the synergistic effect of ISO and insulin on stimulation of PDE3B activity.
PGE1 is one of prostagladins which have an antilipolytic effect (Vassaux et al.,
1992). The precise mechanism by which PGE1 activates PDE remains unknown. The ginseng and PGE2 studies showed that PDE4 partially mediated ginseng and PGE2 antilipolysis. Whether PGE2 and ginseng activate PDE4 in rat adipocytes was investigated in the present study.
6.2 Determination of suitable concentrations of PDE3 and PDE4 inhibitors
Rationale: Four cAMP-hydrolyzing PDEs, VIZ., PDE1, PDE2, PDE3, and PDE4, exist in differentiated 3T3-L1 adipocytes or rat adipocytes (Coudray et al., 1999; Elks and Manganiello, 1984; Shechter, 1984). Therefore, the total PDE activity in rat adipocytes is expected to equal the sum of at least these four PDE activities. When using cilostamide or rolipram to calculate the PDE3 or PDE4 activity in rat adipocytes, it is possible to underestimate the PDE3 or PDE4 activity if the concentration of cilostamide or rolipram is low. On the other hand, when the concentration of cilostamide or rolipram is high, there may be an overlapping inhibition between PDE3 105 and PDE4 and thus PDE3 or PDE4 activity is overestimated. Therefore, selecting the suitable concentrations of cilostamide or rolipram is a critical step for differentiating
PDE3 and PDE4 activity in rat adipocytes.
Hypothesis: If a suitable combination of cilostamide and rolipram is used to detect the activities of PDE3 and PDE4, then the amount detected by the combination of cilostamide and rolipram will be less than the total PDE activity and almost equal to the sum of the activities of PDE3 and PDE4 determined by cilostamide and rolipram alone, respectively.
Experimental design: Three different concentration combinations of cilostamide and rolipram were used to differentiate PDE3 and PDE4 activity in the homogenates: cilostamide at 0.5 µM and rolipram at 5 µM; cilostamide at 0.75 µM and rolipram at
7.5 µM; cilostamide at 1 µM and rolipram at 10 µM. Total PDE activity, PDE3 activity, PDE4 activity, and the combined activity of PDE3 and PDE4 were determined. Detailed information for PDE activity assay and the calculation of PDE isoform activity can be found in Section 2.10.
Results: As shown in Table 6.1, the sum of PDE3 and PDE4 activity in homogenates incubated with 0.5 µM cilostamide and 5 µM rolipram respectively was similar to the amount determined by the combined inhibitors (P>0.9) and was less than the total PDE activity (P<0.03). The sum of PDE3 and PDE4 activity in homogenates incubated with 0.75 µM cilostamide and 7.5 µM rolipram respectively was slightly more than the amount determined by the combined inhibitors (P>0.2) and was less than the total PDE activity (P<0.03). The sum of PDE3 and PDE4 activity in homogenates incubated with 1 µM cilostamide and 10 µM rolipram respectively, was not only more than the amount determined by the combined inhibitors (P>0.1), but
106 also near to the total PDE activity (P>0.2). Therefore, the combination of cilostamide at 0.5 µM and rolipram at 5 µM was suitable for differentiating PDE3 and PDE4 activity in rat adipocytes.
6.3 Subcellular distribution of PDE4 activity in rat adipocytes
Rationale: PDE3B is a membrane-bound protein (Degerman et al., 1998). PDE4 family are both cytosol-soluble and membrane-bound proteins (Houslay, 2001).
PDE4A1 (Pooley et al., 1997) exists exclusively in the particulate fraction; PDE4A4
(Huston et al., 1996), PDE4A5 (McPhee et al., 1995), and PDE4A8 (Bolger et al.,
1996) exist in both the particulate and cytosolic fractions. PDE4B1, PDE4B2, and
PDE4B3 exist in both the cytosolic and the particulate fractions (Huston et al., 1997).
PDE4D1 and PDE4D2 exist only in the cytosolic fraction; PDE4D3, PDE4D4, and
PDE4D5 exist in both the cytosolic and particulate fractions (Bolger et al., 1997). The determination of subcellular distribution of PDE4 activity may help define PDE4 isoform expression in rat adipocytes.
Hypothesis: If PDE4 family are both cytosol-soluble and membrane-bound proteins, then PDE4 activity will be measurable in the particulate fraction and supernatant fraction.
Experimental design: PDE3 and PDE4 activity were differentiated among the whole homogenates (H), the 10,000 ×g pellet fraction (P1), the 100,000 ×g pellet fraction (P2), and the 100,000 ×g supernatant fraction (S) by using 0.5 µM cilostamide and 5 µM rolipram, respectively. Detailed information for PDE activity assay and the calculation of PDE isoform activity can be found in Section 2.10.
Results: As shown in Figure 6.1, the activity of PDE3 and PDE4 in whole homogenates (H) accounted for 65.7% and 19.2% of the total PDE activity, 107 respectively. The activity of PDE3 and PDE4 in the 10,000 ×g pellet fraction (P1), accounted for 79.3% and 13.6% of the total PDE activity, respectively. The activity of
PDE3 and PDE4 in the 100,000 ×g pellet fraction (P2) accounted for 90% and 10.1% of the total PDE activity, respectively. The activity of PDE3 and PDE4 in the 100,000
×g supernatant fraction (S) accounted for 41.1% and 31.9% of the total PDE activity, respectively.
6.4 Effects of PGE2, PIA, and KGE on PDE activity in rat adipocytes
Rationale: ISO and insulin activate particulate PDE3B through different mechanisms in rat adipocytes (Smith and Manganiello, 1989). ISO phosphorylates and activates PDE3B by PKA, whereas insulin phosphorylates and activates PDE3B by PKB (Smith et al., 1991). It is also known that PGE1 activates total PDE in human adipose tissue. The PDE activity assay can detect the changes in PDE activity induced by the above agents. Therefore, PDE activity assay is a suitable approach to determine if PGE2 and ginseng activate PDE.
Hypothesis: If PGE2 and ginseng antilipolysis is reduced by a specific PDE4 inhibitor, then PGE2 and ginseng will activate PDE4 in rat adipocytes. If PIA antilipolysis is not reduced by a specific PDE4 inhibitor, then PIA will not activate
PDE4 in rat adipocytes.
Experimental design: All reagents for treating adipocytes were prepared just before incubations to ensure stability. Freshly isolated adipocytes were incubated with
ISO (100 nM), insulin (90 nM), PGE2 (1 µM), PIA (100 nM), and KGE (100 µg/ml) for 15 min. The particulate fraction (100,000 × g pellet) of rat adipocyte was used in this assay. ISO and insulin were used as positive controls in this assay. In the lipolysis assay, PDE isoform inhibitors (cilostamide at 5 µM and rolipram at 10 µM) were 108 incubated with adipocytes for 1 hour. In contrast with the lipolysis assay, PDE isoform inhibitors (cilostamide at 0.5 µM and rolipram at 5 µM) were incubated with the particulate fraction for 10 min in PDE activity assay. The rationale for these changes on the incubation concentration and time is that PDE isoform inhibitors work directly on adipocyte extracts rather than intact adipocytes. Detailed information on
PDE activity assay can be found in Section 2.10.
Results: The effects of various agents on the total PDE, PDE3, and PDE4 activity in isolated adipocytes are shown in Figure 6.2. ISO, insulin, and PGE2 increased the total PDE activity by 65.2% (p<0.01), 30.6% (p<0.01), and 15.7%, respectively
(p<0.01). PIA inhibited the total PDE activity by 17.1% (p<0.01). Ginseng did not change the total PDE activity. ISO, insulin, and PGE2 increased PDE3 activity by
86.3% (p<0.01), 24.3% (p<0.05), and 21.4% (p<0.01), respectively. These agents also stimulated PDE4 activity by 108.4%, 23.6%, and 26.7%, respectively, although effects were not statistically significant (Table 6.2).
6.5 Discussion
In the present study, the activation of PDE activity in rat adipocytes was investigated. Lipolysis assay is an end-point assay and PDE activity assay is a more direct assay to detect the activation of PDE. The results of these two assays are complimentary on addressing the role of PDE in rat adipocytes.
A previous study with rat adipocytes showed that PDE3 determined by OPC3911
(3 µM) accounted for more than 90% of total PDE activity in the particulate fraction
(50,000 ×g) and about 45% of that in the supernatant fraction (50,000 ×g) assayed at the concentration of 0.5 µM cAMP (Eriksson et al., 1995). Consistent with this study,
PDE3 activity as determined using cilostamide (5 µM) in the present study accounted 109 for 79.3% and 90% of the total PDE activity in the 10,000 ×g pellet fraction and the
100,000 ×g pellet fraction, respectively, when the concentration of cAMP was 0.1 µM.
Also, PDE3 activity represented 41% of total PDE activity in the 100,000 ×g supernatant fraction.
The above study also showed that PDE4 activity accounted for 10% of total PDE in whole rat adipocytes as determined in the presence of Ro 20-1724 (30 µM)
(Eriksson et al., 1995). PDE4 activity as determined in the presence of rolipram (10
µM) in the present study accounted for 19% of the total PDE activity in whole homogenates. The discrepancy likely results from the use of different PDE4 inhibitors.
Furthermore, the present study demonstrated that PDE4 activity accounted for 14%,
10%, and 32% of the total PDE activity in the 10,000 ×g pellet fraction, the 100,000
×g pellet fraction, and the 100,000 ×g supernatant fraction, respectively. These results demonstrate that the absolute PDE4 activity was almost equivalent among three subcellular fractions, and that the proportion of PDE4 activity relative to the total
PDE was the highest in the supernatant fraction. It is unknown if this distribution of
PDE4 increases the efficiency for cAMP degradation in rat adipocytes. Based on the subcellular distribution of PDE4 in the present study, it is not possible to deduce which PDE4 isoform is expressed in adipocytes.
In agreement with previous studies (Engfeldt et al., 1983; Smith and
Manganiello, 1989), ISO and insulin increased the total PDE and PDE3 activity in the present study. In a previous study, adenosine (70 µM) inhibited PDE activity by 31%,
19%, and 10% in the supernatant (48,000 ×g) of rat adipocyte homogenates assayed at the concentrations of 0.1 µM, 0.2 µM, and 0.5 µM cAMP, respectively (Fain et al.,
110 1972). In the present study, PIA (100 nM) inhibited the particulate total PDE activity by 17%. The mechanism by which adenosine inhibits PDE activity in rat adipocytes is not clear.
Result with ginseng did not support the hypothesis. It is possible that the activation of PDE in rat adipocytes stimulated by 100 µg/ml ginseng was not sufficient to detect the change in activity. According to the previous study (Smith et al., 1991), the activation signal of PDE3B in rat adipocytes induced by insulin and
ISO was much less than the phosphorylation signal (3-fold vs. 40-fold). Therefore, the concentrations of insulin and PGE2 used in the PDE assay were much higher than in the lipolysis assay to obtain maximal stimulation. Since ginseng is a complex mixture of compounds, the concentration of ginseng in the PDE assay was not increased to prevent non-specific effects.
In the present study, PGE2 increased the total PDE activity, PDE3 activity, and
PDE4 activity by 16%, 21%, and 27%, respectively. In the lipolysis assay, PGE2 antilipolysis was reduced only by a PDE4 inhibitor, but not by a PDE3 inhibitor. This inconsistency may result from the different experimental systems. Lipolysis assay was performed using intact adipocytes, whereas PDE activity assay was performed using adipocyte extracts.
With the availability of the cAMP biosensor in recent years, the diffusion of cAMP in living cells can be tracked (Zaccolo and Pozzan, 2002). It has been known that cAMP is compartmentalized in a variety of cell types because the activity of phosphodiesterases limits free diffusion of this second messenger (Zaccolo and
Pozzan, 2002). It is possible that only PDE4 is accessible to the endogenous cAMP pool in cytosol even if both PDE3 and PDE4 are activated by PGE2 in rat adipocytes.
This situation would limit the role of PDE3 in PGE2 antilipolysis. 111 The mechanisms by which PGE2 up-regulates PDE4 activity in rat adipocytes are not clear. A model was proposed to explain how ERK activation affects the activity of the long PDE4D5 isoform in human aortic smooth-muscle cells (Baillie et al., 2001).
In this model, ERK activation initially inhibits the activity of PDE4D5. Meanwhile,
ERK activation results in the generation of PGE2 through the metabolism of arachidonic acid. PGE2 stimulates adenylate cyclase activity by binding to the EP2 receptor, leading to an increase in cAMP concentration. Subsequently, activation of
PKA results in the phosphorylation and activation of PDE4D5. The activation of
PDE4D5 in ERK-dependent manner counteracted the ERK-mediated inhibition of
PDE4D5, so the net effect is activation of PDE4D5. However, this model is not applicable for PGE2-mediated PDE4 activation in rat adipocytes. It has been known that EP1, EP3 and EP4 are expressed in mature adipocytes. Among them, only EP3 is expressed exclusively in mature adipocytes (Borglum et al., 1999). PGE2 inhibits adenylate cyclase activity by binding to EP3, while it stimulates adenylate cyclase by binding to EP4 (Narumiya et al., 1999). Currently, it is not known whether the binding affinity of these two PGE2 receptors is different in rat adipocytes. If PGE2 binds to both EP3 and EP4, the activity of adenylate cyclase will be regulated by Gs and Gi differentially. Because the net effect of PGE2 stimulation in rat adipocytes is the inhibition of adenylate cyclase, it is possible that the expression of EP3 or binding to
EP3 is predominant in rat adipocytes.
In summary, the present study showed PGE2 stimulated the total PDE, PDE3, and PDE4 in rat adipocytes. This finding is in accordance with the result of PGE2 lipolysis assay. Although PDE3 and PDE4 also mediated ginseng antilipolysis, stimulation of PDE with ginseng was not observed in the present study.
112
Activity (pmol cAMP/min/mg protein)
Total PDE3 PDE4 Sum of PDE3+4 PDE (a) (b) (a)+(b) CIL (0.5 µM) + ROL (5 µM) 54.7±7.9 35.3±7.3 10.1±3.2 45.4±9.3* 45.7±7.2
CIL (0.75 µM)+ROL (7.5 µM) 54.7±7.9 39.2±7.1 10.6±2.9 49.8±8.6* 47.1±7.5
CIL (1 µM)+ROL (10 µM) 54.7±7.9 39.9±6.9 13.1±2.5 53.0±7.6 48.7±7.5
Table 6.1. Effects of different concentrations of the PDE3 and PDE4 inhibitors on PDE3 and PDE4 activity in rat adipocytes. The total PDE, PDE3, and PDE4 were determined in the absence and presence of cilostamide (CIL), rolipram (ROL), and the combination of two inhibitors at the respective concentrations as indicated above. The sum of PDE3 (a) and PDE4 (b) activity in homogenates incubated with 0.5 µM cilostamide and 5 µM rolipram respectively almost equaled the amount determined by the combination of the two inhibitors and was less than the total PDE activity. PDE isoform activity was calculated as the difference in activity in the absence and presence of inhibitor(s). Total PDE is the activity for samples incubated without inhibitors. Data are mean ± SEM of 3 independent experiments assayed in triplicate. Statistical significance compared to total PDE activity is denoted with *, P <0.05.
113
T-PDE PDE3 PDE4
120 100
80
60
40 20
0 pmol cAMP/min/mg protein HP1P2S
Figure 6.1. Subcellular distribution of total PDE, PDE3B, and PDE4 activity in rat adipocytes assayed with 0.1 µM cAMP. PDE4 activity accounted for 19.2% of total PDE activity in homogenates (H). PDE4 activity was evenly distributed across the 10,000 × g pellet fraction (P1), the 100,000 × g pellet fraction (P2), and the 100,000 × g supernatant fraction (S). The proportion of PDE4 activity in the 100,000 × g supernatant fraction is the highest among three subcellular fractions. Data are mean ± SEM of 5 independent experiments assayed in triplicate. H, homogenate; P1, 10,000 × g pellet fraction; P2, 100,000 × g pellet fraction; S, 100,000 × g supernatant fraction.
114
120 * 100 * 80 * 60 * 40 20 0
pmol cAMP/min/mg protein ) ) ) ) l) M M M M n m Basal 0 n 0 0 n 0 (1 u 0 (9 (1 s o s In GE2 (100 ug/ I P PIA (1 KGE
Figure 6.2. Effects of various agents on total PDE activity in rat adipocytes. Isolated rat adipocytes were incubated with agents indicated above for 15 min. The total PDE activity in the particulate pellet fraction was assayed at 0.1 µM cAMP. Basal indicates that no treatment was present. Data are mean ± SEM of 5 independent experiments assayed in triplicate. Statistical significance compared to basal is denoted with *, P <0.05.
115
Activity (pmol cAMP/min/mg protein)
PDE3 PDE4
Basal 48.8±5.6 12.1±2.2
Iso (100 nM) 90.9±11.8* 25.3±4.5
Ins (90 nM) 60.6±5.6* 15.0±2.7
PGE2 (1 µM) 59.2±6.5* 15.4±3.0
Table 6.2. Effects of various agents on PDE3 and PDE4 activity in rat adipocytes. Isolated adipocytes were incubated with various agents as indicated above for 15 min. PDE3 and PDE4 activity in the particulate pellet fraction was assayed in presence of 0.1 µM cAMP. PDE3 activity was calculated as the difference between the total activity and the activity inhibited by 0.5 µM cilostamide; PDE4 activity was calculated as the difference between the total activity and the activity inhibited by 5 µM rolipram. Basal indicates that no treatment was present. Data are mean ± SEM of 5 independent experiments assayed in triplicate. Statistical significance compared to basal is denoted with *, P <0.05.
116
CHAPTER 7
CONCLUSIONS
It is generally thought that FFA play a key role in the development of insulin resistance and type 2 diabetes. Increased circulating FFA concentration and concomitant accumulation of lipid in nonadipose organs and tissues are common in obese people and people with type 2 diabetes. Adipose tissue is considered as the primary site for insulin resistance. FFA-induced insulin resistance decreases glucose utilization by skeletal muscle by inhibiting glucose uptake and glycogen synthesis, stimulates hepatic glucose production by increasing gluconeogenesis and glycogenolysis, and reduces insulin extraction by the liver. Furthermore, long-term exposure of pancreatic β-cells to high level of FFA impairs the glucose-stimulated insulin secretion, and lipid accumulation in β cells leads to β-cell apoptosis.
The high circulating FFA level in insulin resistance and type 2 diabetes results from an increase in rate of lipolysis. Lipolysis is regulated by the intracellular concentration of cAMP. Many endogenous and exogenous factors affect the rate of lipolysis by regulating cAMP concentration. Catecholamines increase cAMP concentration by activating adenylate cyclase; insulin enhances cAMP degradation by activating PDE3B; PGE2 and adenosine decrease cAMP concentration by inhibiting adenylate cyclase. It is thought that PGE2 and adenosine are endogenous inhibitors of lipolysis and play a tonic role in the inhibition of lipolysis in the physiological state.
117 It is well established that PDE3B is the major isoform of PDE in rat adipocytes.
Insulin exerts its antilipolytic effect by phosphorylating and activating PDE3B. In addition to PDE3B, PDE4 activity is also detected in rat adipocytes by biochemical and pharmacological methods. In contrast with PDE3B, PDE4 does not mediate the antilipolytic effect of insulin. Therefore, the expression and the physiological role of
PDE4 in rat adipocytes remain unclear. In the present study, the PDE4 expression in rat adipocytes was characterized at the mRNA, protein and enzymatic activity levels.
RT-PCR and QPCR showed that all four PDE4 subtypes are expressed in rat adipocytes and the expression of four PDE4 subtypes (A, B, C, D) relative to PDE3B was 7%, 19%, 19%, and 7%, respectively. Western blot showed that PDE4B3 (92 kDa) might exist in the cytosol fraction of rat adipocytes. PDE4 activity accounted for 19% of the total PDE activity in the homogenates of rat adipocytes. Moreover, the specific
PDE3 inhibitor cilostamide and the specific PDE4 inhibitor rolipram increased basal lipolysis by 28% and 33%. These data indicated that all four PDE4 subtypes were expressed in rat adipocytes and PDE4 played a role in the regulation of basal lipolysis.
The present study showed that ginseng has an antilipolytic effect in rat adipocytes. Ginseng antilipolysis was mediated by both PDE3 and PDE4. The signaling pathway for ginseng antilipolysis was different from that activated by insulin. PGE2 and adenosine antilipolysis were also compared with ginseng antilipolysis. There were some similarities and divergences among these three kinds of antilipolysis. The PDE4 specific inhibitor rolipram alone reduced ginseng and
PGE2 antilipolysis, but not adenosine antilipolysis. Combination of the PDE3 and
PDE4 inhibitors had a synergistic effect on PGE2 and adenosine antilipolysis, and had an additive effect on ginseng antilipolysis. It can be inferred that ginseng and PGE2 118 antilipolysis were mediated in part by PDE4. The synergistic effect of two inhibitors may result from the increase in the concentration of cAMP by inhibiting both PDE3 and PDE4.
It is generally accepted that PGE2 inhibits lipolysis by inhibiting adenylate cyclase and thus decreasing the intracellular cAMP concentration. The results of the present study suggest that there are at least two mechanisms underlying the antilipolytic effect of PGE2. In addition to the inhibition of adenylate cyclase, activation of PDE4 also contributes to PGE2 antilipolysis. The results of PDE activity assay confirmed that PGE2 slightly increased the total PDE activity, PDE3 activity, and PDE4 activity.
In summary, the present study demonstrated that all four PDE4 subtypes were expressed in rat adipocytes and PGE2 inhibited lipolysis by activating PDE4 in part.
Ginseng mimics the antilipolytic effect of insulin, but the signaling pathway differes for these two effects. Ginseng antilipolysis was also mediated by PDE4 in part.
119
Epilogue
The present study focused on the expression and the antilipolytic role of PDE4 in rat adipocytes. There are several areas related to PDE4-mediated antilipolysis that require future investigation.
First, the expression of PDE4 subtypes in human adipocytes needs to be determined. To our knowledge, there is no available information on the expression of
PDE4 in human adipocytes. It is possible that the expression of PDE4 is different from that of rat adipocytes because of species specificity.
Second, additional investigation is necessary to identify the active components for ginseng antilipolysis. It is known that six major ginsenosides are not responsible for ginseng antilipolysis individually. Besides adenosine, Korean ginseng contains another antilipolytic agent, pyro-glutamate. It is not known whether the antilipolytic effect of pyro-glutamate can be reversed by the PDE3 or PDE4 inhibitors.
Third, further work is needed to investigate the mechanisms by which PGE2 activates PDE in adipocytes. Because the increase in PDE activity stimulated by PGE2 occurred quickly, it can be inferred that it is a post-translational regulation, e.g. phosphorylation. Little is known about the signaling pathway to activate PDE by
PGE2 stimulation in rat adipocytes.
Finally, in vivo study is needed to further evaluate the antilipolytic effect of ginseng, especially on obese or diabetic animal models. Since ginseng slightly stimulates glucose transport into adipocytes in vitro, it is unclear if inhibition of lipolysis is the major mechanism responsible for the hypoglycemic effect of ginseng. 120
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