UNIVERSITY OF CINCINNATI

Date:______

I, ______, hereby submit this work as part of the requirements for the degree of: in:

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Metabolic Functions of Prolactin in the Mouse

A dissertation submitted to the division of graduate studies and research of the University of Cincinnati

In partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

in the Neuroscience Graduate Program of the College of Medicine

October 2006

By

Christopher R. LaPensee

Bachelor of Science, University of Michigan, 2000

Committee Chair: Nira Ben-Jonathan, Ph.D. Scott Belcher, Ph.D. Nelson Horseman, Ph.D. Randy Seeley, Ph. D. Patrick Tso, Ph.D. Abstract

PRL is a 23 kDa hormone produced primarily in the pituitary that is best recognized for its lactogenic activity. However, PRL is also involved in the regulation of metabolic homeostasis in many species. Studies in rats generally indicate that PRL promotes weight gain, while in humans, the association between PRL and adiposity is unclear. Several mouse models and adipocyte cell lines have been used to study the effects of PRL on growth, adipokine secretion, and lipid and glucose metabolism, but a consensus has not emerged, given limited and often conflicting data. Thus, clarifying the metabolic functions of PRL in the mouse would address many published controversies and contribute to understanding the systems and factors that regulate energy balance.

Our hypothesis was that PRL is required for normal weight gain and adiposity in male and non- lactating female mice. We postulated that this may occur by PRL-mediated alterations in adipokine secretion, glucose metabolism, and/or adipose tissue growth and metabolism. To assess the metabolic role of PRL in vivo, we first compared 1) weight gain, 2) body composition, 3) serum lipid profile, 4) circulating leptin and adiponectin levels, and 5) glucose tolerance in PRL-knockout, heterozygous, and wild-type mice maintained on high-fat, low-fat, and standard chow. We found that PRL-deficiency had no effect on the rate of weight gain, body composition, serum lipids, or adiponectin levels in either sex on any diet. Glucose tolerance was slightly impaired in 4-week old male pups, but not in adults or in females at any age. Leptin was elevated in male, but not female PRL-deficient mice on a low-fat diet.

We then measured the expression of selected genes in the adipose tissue and liver of PRL- knockout and wild-type mice under fed and fasted conditions. Nearly all genes, including lipoprotein lipase, hormone-sensitive lipase, leptin, adiponectin, and resistin, were unchanged in PRL-deficient mice when compared to wild-type mice. One novel factor, fasting-induced adipocyte factor (FIAF) was upregulated by fasting in both adipose tissue and liver of wild-type mice, but not in male PRL-deficient mice. When serum from these animals was analyzed, two isoforms of FIAF were significantly higher in PRL-deficient male mice than wild-types.

To determine the effects of PRL on adipocyte metabolism, we utilized the murine 3T3-L1 cell line, as well as cultured adipose tissue explants. PRL at physiological doses had no effect on preadipocyte proliferation, lipid accumulation, or morphological alterations during differentiation in 3T3-L1 cells. Chronic PRL treatment had inconsistent effects on basal and insulin-stimulated glucose uptake, as well as on glucose transporter 4 (GLUT4) expression in mature 3T3-L1 adipocytes, while short-term PRL treatment had no effect on these parameters. Finally, PRL did not affect basal or stimulated lipolysis in adipose tissue explants from mice, but significantly inhibited glycerol release from both rat and human adipose tissue explants in a dose-dependent manner.

In conclusion, PRL likely plays a role in regulating adipokine release and glucose metabolism, but does not affect gross metabolic parameters in mice. Adiposity is a reflection of adipocyte growth and lipid storage, both of which are unaffected by PRL treatment in vitro. The lipolytic actions of PRL vary among species, with no effects observed in mice. Thus, the metabolic functions of PRL in the mouse are limited compared to other species.

Acknowledgements

I would like to extend sincere thanks to my advisor, Dr. Nira Ben-Jonathan. Your guidance ensured that I remained focused and strived to do my best. Your door was always open for a conversation, and you carefully listened to and considered my opinion. I will always value the way in which you taught me think critically and write effectively. Thank you.

To my thesis committee members, thank you for your insightful observations and helpful suggestions. You allowed me to explore numerous aspects of metabolism, which allowed me to learn a significant number of techniques and gain a wealth of knowledge. Dr. Horseman, thank you for giving me the opportunity to study the PRL-deficient mouse.

To the Ben-Jonathan laboratory, thank you for being an easy group to work with, and for always being accommodating and helpful. It was important that we all share ideas and techniques as we in ventured together into a new field, and you were all excellent at doing so.

I would like to thank the Neuroscience Graduate Program and the Department of Cell and Cancer Biology for their support. I am grateful to all of the students and faculty that I have interacted with over the years.

Of course, I will forever cherish the unconditional love and support that I received from my family. Having my mother’s, father’s, and grandparent’s support over the years has given me the confidence and ability to pursue a career in science. I truly thank you for all you have given me. I love you.

Lastly, to my wife Beth, you have made my life complete. You are my best friend and have brought me happiness that I could have never imagined. As a fellow student, you understand the ups and downs of graduate school and scientific research, and you always keep me thinking positive. Thank you for being a part of my life. I love you.

Table of Contents

Table of Contents ...... 1 Figures and Tables...... 3 Abbreviations ...... 5 Introduction...... 9 Chapter 1. Literature Review ...... 11 Section I. Key tissues involved in metabolism ...... 12 A. Adipose Tissue...... 12 1. Depots and cell types ...... 12 2. Adipogenesis...... 14 3. Lipid metabolism ...... 18 Lipogenesis ...... 18 Fatty Acid Synthase (FAS) ...... 21 Hormone Sensitive Lipase (HSL)...... 22 Perilipin...... 24 4. Glucose Transport...... 26 2. Endocrine function...... 30 Leptin ...... 30 Adiponectin...... 33 Fasting-induced adipocyte factor (FIAF)...... 35 B. Pancreas ...... 37 1. Structure...... 37 2. β cells and insulin synthesis...... 38 3. Insulin secretion and the role of glucose...... 39 C. Liver...... 41 1. Glucose metabolism...... 41 2. Lipid metabolism ...... 43 D. Brain...... 44 Section II. Integrative regulation of metabolic homeostasis...... 48 A. Overall Considerations...... 48 B. Energy storage and utilization ...... 50 C. Hormonal signals ...... 52 Section III. Prolactin ...... 55 A. Gene structure and regulation of expression...... 55 B. Protein structure and variants...... 56 C. Pituitary lactotrophs...... 57 D. Regulation of pituitary PRL secretion ...... 59 E. Physiology of PRL...... 62 Section V. Prolactin Receptors...... 64 A. Protein Structure ...... 64 B. Gene structure ...... 65 C. Activation and signal transduction...... 66 D. Receptor distribution and regulation of expression ...... 69 Section VI. Biological Functions of PRL...... 70 A. Mammary gland development ...... 70

1 B. Luteal Function ...... 72 C. Osmoregulation...... 72 D. Immunoregulation...... 73 E. Behavior...... 74 F. Prostate...... 75 Section VI. Involvement of PRL in metabolic homeostasis...... 75 A. Body weight, adiposity, and food intake ...... 75 B. Lactogenesis...... 78 C. Adipokine release...... 80 D. Lipid Metabolism...... 81 E. Pancreas ...... 84 F. Liver...... 86 G. Prostate...... 87 References...... 89 Chapter 2. Specific Aims of Thesis Research ...... 128 Chapter 3. The PRL-Deficient Mouse has an Unaltered Metabolic Phenotype ...... 131 Abstract...... 133 Introduction...... 134 Materials and methods ...... 135 Results...... 137 Discussion...... 139 Acknowledgements...... 142 References...... 142 Figure Legends...... 146 Chapter 4. Expression of Selected Genes in PRL-Deficient Mice ...... 155 Introduction...... 156 Materials and methods ...... 156 Results...... 157 Discussion...... 158 References...... 160 Chapter 5. The Role of PRL in Preadipocyte Proliferation and Lipid Accumulation...... 168 Introduction...... 169 Materials and methods ...... 170 Results...... 171 Discussion...... 173 References...... 176 Chapter 6. The Effects of PRL on Glucose Uptake in Mature Adipocytes ...... 185 Introduction...... 186 Materials and methods ...... 186 Results...... 187 Discussion...... 188 References...... 190 Chapter 7. General Conclusions...... 195 Chapter 8. Curriculum Vitae...... 200

2 Figures and Tables

Chapter 1. Literature review Figure 1. Overview of adipogenesis…………………………………………………...... 15 Figure 2. Lipid flux in adipocytes……………………………………………………...... 20 Figure 3. Schematic of glucose uptake…………………………………………………..…. ..29 Figure 4. Integrated metabolic homeostasis……………………………………………...... 53 Figure 5. The PRL gene and protein……………………………………………………...... 57 Figure 6. The mouse PRLR………………………………………………………………….. 65 Figure 7. PRLR signaling……………………………………………………………………. 68 Figure 8. Putative functions of PRL in adipocytes………………………………………...... 82

Chapter 3. Metabolic characterization of the PRL-deficient mouse Figure 1. PRL and PRLR levels in wild-type, heterozygous, and PRL-knockout mice….... 148 Figure 2. Growth curve……………………………………………………………………...149 Figure 3. Body composition………………………………………………………………... 150 Figure 4. Serum lipids……………………………………………………………………….151 Figure 5. Serum leptin and adiponectin concentrations……………………………………. 152 Figure 6. Glucose tolerance determination…………………………………………………. 153 Figure 7. Effects of PRL lipolysis in adipose tissue explants……………………………….154

Chapter 4. Expression of selected genes in PRL-knockout mice Table 1. PCR primer sequences…………………………………………………………… 162 Table 2. Gene expression in adipose tissue and liver………………………………………163 Figure 1. FIAF gene expression in males………………………………………………….. 164 Figure 2. Effect of fasting on leptin and LPL gene expression…………………………….. 165 Figure 3. Serum FIAF levels in male mice………………………………………………….166 Figure 4. Quantitative measurement of serum FIAF levels in male mice………………….. 167

Chapter 5. The role of PRL in preadipocyte proliferation and lipid accumulation Figure 1. 3T3-L1 preadipocyte viability plot………………………………………………. 178 Figure 2. 3T3-L1 standard growth curve…………………………………………………… 179 Figure 3. Effect of PRL on preadipocyte viability…………………………………………. 180 Figure 4. Effect of PRL on preadipocyte viability…………………………………………. 181 Figure 5. PRL and PRLR expression during 3T3-L1 differentiation………………………. 182 Figure 6. Effect of PRL on lipid accumulation during adipogenesis………………………. 183 Figure 7. Effect of PRL on morphological changes during adipogenesis………………….. 184

Chapter 6. The effects of PRL on glucose uptake in mature adipocytes Figure 1. Effect of chronic PRL treatment on glucose uptake……………………………... 192 Figure 2. Effect of short-term PRL treatment on glucose uptake…………………………...193 Figure 3. GLUT4 gene expression in PRL treated adipocytes……………………………... 194

3 4 Abbreviations

-/- gene deficient +/- heterozygous +/+ wild-type aa amino acids ACC acetyl-CoA carboxylase ACP acyl carrier protein ACRP adiponectin AdipoR adiponectin receptor AOX acyl CoA oxidase aP2 adipose promoter 2 Asp aspartate ADP adenosine diphosphate AgRP agouti-related protein AMP adenosine monophosphate AMPK adenosine monophosphate kinase AP anterior pituitary aPKC atypical protein kinase C apo apoliprotein APS adaptor molecule containing pleckstrin homology and SH2 domains ATP adenosine triphosphate ATGL adipose triglyceride lipase BAT brown adipose tissue BMI body mass index Ca calcium CAM cell adhesion molecule cAMP cyclic adenosine monophosphate CAP cb1-associated protein CART cocaine and amphetamine-regulated transcript cDNA complementary deoxyribonucleic acid C/EBP CCAAT-enhancer binding protein CNS central nervous system CoA coenzyme A CRE carbohydrate response element CRH corticotrophin-releasing hormone CSF cerebrospinal fluid D2R dopamine receptor-2 DA dopamine db diabetic DG diglyceride DMH dorsomedial hypothalamus DNA deoxyribonucleic acid ECM extracellular matrix EGF epidermal growth factor ER estrogen receptor

5 ERC endosomal recycling compartment FAS fatty acid synthase FBS fetal bovine serum FFA free fatty acid FIAF fasting-induced adipocyte factor GAS gamma interferon activation site GH growth hormone GHRH growth hormone−releasing hormone GK glucokinase GLUT glucose transporter GSV glucose transporter storage vesicle GTP guanosine triphosphate HDL high density lipoprotein His histodine HMW high–molecular weight HPA hypothalamic-pituitary-adrenal I- intermediate ICDH isocitrate dehydrogenase i.c.v. Intracerebroventricular IDL intermediate density lipoproteins IGF insulin-like growth factor IL interleukin IP intermediate pituitary IR insulin receptor IRE insulin response element IRS insulin receptor substrate kDa kilo Dalton Jak janus kinase L- long LCFA long chain fatty acid LD lipid droplet LDL low density lipoprotein LH luteinizing hormone LHA lateral hypothalamic area LMW low–molecular weight LPL lipoprotein lipase mAAT mitochondrial aspartate aminotransferase MAPK mitogen activated protein kinase MAT malonyl-CoA-/acetyl-CoA-ACP-transacylase MC melanocortin receptor MCH melanin-concentrating hormone ME median eminance MG monoglyceride MGL monoglyceride lipase MMW middle–molecular weight mPOA medial preoptic area

6 mRNA messenger ribonucleic acid MSH melanin-stimulating hormone NADPH nicotinamide adenine dinucleotide phosphate-oxidase NEFA non-esterified fatty acid NL neural lobe NPY Neuropeptide Y OA oleic acid Ob obesity PCR polymerase chain reaction PDK1 3-phosphoinositide-dependent protein kinase-1 PH pleckstrin homology Pi inorganic phosphate PI3K phosphotidylinositol-3-kinase PIKfyve phosphoinositide 5-kinase containing a Fyve finger PIP2 phosphatidylinositol 4,5-bisphosphate PIP3 phosphatidylinositol 3,4,5-trisphosphate Pit-1 pituitary-specific transcription factor PKA protein kinase A PKC protein kinase C PL placental lactogen POMC proopiomelanocortin PP-1 protein phosphatase-1 PP posterior pituitary PPAR peroxisome proliferator-activated receptor PPRE peroxisome proliferator response element PPY pancreatice peptide Y Pref-1 preadipocyte expression factor 1 PRL prolactin PRLR prolactin receptor PVN paraventricular nucleus R receptor RIA radioimmunoassay RNA ribonucleic acid RT reverse transcription RXR retinoic acid receptor S- short SCD1 stearoyl-CoA desaturase Ser serine SH2 src homology 2 SNARE soluble N-ethylmaleimide sensitive factor-attachment receptor SOCS suppressor of cytokine signaling Sp stimulatory protein SREBP sterol regulatory binding protein Stat signal transducer and activator of transcription TG triglyceride TGF transforming growth factor

7 THDA tuberohypophysial dopaminergic TIDA tuberoinfundibular dopaminergic TNF tumor necrosis factor Trp tryptophan TRH thryotropin releasing hormone TSH thyroid-stimulating hormone TU tuberal hypothalamus TZD thiazolidinediones UCP-1 uncoupling protein 1 UDP uridine diphosphate UST upstream stimulatory factor VAMP2 vesicle-associated membrane protein 2 VIP Vasoactive intestinal peptide VLDL very low density lipoprotein VMN ventromedial nucleus of the hypothalamus WAP whey acidic protein WAT white adipose tissue

8 Introduction

Metabolism is the sum of all of the chemical reactions an organism needs to survive. It is the principal and indispensable process that supports life-long homeostasis. The process of metabolism is a balancing act involving two activities that are constantly taking place - the anabolic building up of body tissues and energy stores for use in the future, and the catabolic breaking down of large molecules, mostly carbohydrates and fats, to produce the energy required for all activity in cells.

Proper regulation of metabolic homeostasis is crucial for survival. In adult animals and humans, body weight remains within a relatively narrow range, despite large day-to-day fluctuations in the amount of food consumed. This occurs because energy flux is carefully monitored and adjusted to ensure that fuel intake and energy expenditure remain within acceptable limits. Although major changes of body adiposity can be induced in humans and animals by restricting energy intake or by overfeeding, body weight and adiposity return very close to baseline levels when ad libitum feeding is resumed.

Efficient storage and use of energy involves a number of specific pathways whose components are expressed and activated depending upon the energy needs of the organism and physiological activity. The major peripheral organs participating in the coordinated regulation of metabolism include the pancreas, liver, adipose tissue, and skeletal muscle. Each tissue performs specific functions to ensure proper utilization and mobilization of carbohydrates and lipids. Together with the brain, these tissues orchestrate the transition between widely varying states, including the fed state, fasting, and exercise.

Peripheral signals released from these and other tissues reflect short- and long-term nutritional status, and influence central circuits in the hypothalamus and brain stem to modulate neuropeptide release and hence food intake and energy expenditure. Moreover, these factors can act in an autocrine/paracrine fashion, influencing the metabolic activity of other peripheral tissues, as well as those from which they were secreted. These signals exist in many forms, including hormones and cytokines, as well as circulating energy-rich substrates like fatty acids, glucose, and amino acids.

Although this physiological system is designed to maintain energy balance, complex interactions between genetic and environmental factors can impinge on both peripheral signals and central pathways, and may result in chronic disease states such as obesity, diabetes, and associated disorders. Obesity now affects as many as 30% of the adult population of the United States, and is rapidly on the rise in many other countries. This modern disease has reached epidemic proportions and produces a heavy financial burden on health care systems (1). In large part, the shift towards positive energy balance reflects both increases in energy intake and decreases in physical activity. Moreover, the rising proportion of energy derived from fat is linked to weight gain and obesity. A detailed understanding of the pathogenesis of obesity and its associated complications may make successful treatment possible.

There has been considerable interest in energy balance throughout history. Perhaps the first quantification of metabolic parameters came from Santorio Sanctorius in the 1500s, who slept,

9 ate, worked, and made love in an elaborate weighing chair for thirty years to record how much his weight changed as he ate, fasted, or excreted (2). Often depriving himself of food and drink, Santorio determined that the daily change in body mass approached 1.25 kg. In the early twentieth century Harris and Benedict developed prediction equations for basal energy expenditure from the results of several elaborate studies (3). The equations were developed as normal standards against which basal energy expenditure during disease states could be compared. The equations continue to be used in that capacity, and are also used clinically as a basis for prescribing energy intake for hospitalized patients, and to formulate dietary intake goals for weight loss.

While the study of metabolic homeostasis in humans has advanced in our understanding of energy balance, certain techniques such as brain lesions and gene ablation cannot be performed in humans, necessitating animal and cell models. Indeed, the development of transgenic animals overexpressing or lacking specific genes has proven invaluable in the study of metabolism. Even before the advent of transgenics, animals with spontaneous mutations causing obesity led to the discovery of leptin, probably the most-well known adipokine, and its receptor. However, there are limitations to animal studies. For example, we cannot simply ask animals questions such as “are you hungry”, or “how does that taste?” Due to the personal nature of satiety, the objective study of satiety in non-human animals must therefore measure and draw conclusions from quantifiable behaviors. Additionally, although transgenics are invaluable tools in the study of various physiological systems, a desired or expected phenotype is not always observed. As it pertains to this thesis, this may be particularly true in the study of metabolic homeostasis, which is complicated by the number of pathways involved as well as the complex interplay between interconnected, compensatory and/or redundant systems.

While many hormones such as leptin and insulin are known primarily for their role in metabolic homeostasis, PRL is historically associated with its actions in the mammary gland, where it is a crucial promoter of gland development as well as lactogenesis. However, PRL is a member of the cytokine family, a diverse group of molecules that exert pleiotropic functions, including growth, differentiation, and inflammatory responses. In addition to leptin, other members of this family are also known to regulate several aspects of metabolic homeostasis, including food intake and lipid metabolism, and there is considerable evidence that PRL mediates several related processes in mice. However, as the body of literature pertaining to PRL as a metabolic hormone has grown over the past few decades, its function and importance within this field have come into question.

Our interest in PRL as a metabolic hormone began in 2001, when a study using the PRLR- deficient mouse reported a significant reduction in the rate of weight gain, as well as a marked decrease in abdominal fat mass and serum leptin levels (4). These intriguing observations data strongly implicated PRL in metabolic homeostasis in male and non-lactating female mice. However, studies from other laboratories failed to find a clear relationship between PRL, weight gain, adiposity, and several other metabolic parameters in mice. The availability of PRL- knockout mice presented us with a unique opportunity to study the metabolic actions of PRL in vivo. Additionally, the availability of a murine adipocyte cell line allowed us to investigate several PRL-mediated aspects of metabolism in vitro.

10

Chapter 1. Literature Review

11 Section I. Key tissues involved in metabolism

To study how metabolic homeostasis is so precisely regulated, it is important to understand the organs involved, including the cues they respond to and the specific functions that they perform. Of course, this section can not cover all tissues that participate in the regulation of metabolic homeostasis. Therefore, it is the goal of this section to introduce the major players, with an emphasis on those related to this thesis. The information presented in this section should give the reader a better appreciation for the complex inter-organ fluxes of carbohydrate and lipid-based energy sources.

A. Adipose Tissue

Adipose tissue is a specialized connective tissue that functions primarily to store energy in the form of fat. However, it also serves many other purposes, including heat production, insulation, cushioning, and as an endocrine regulator of homeostasis. This chapter will discuss adipose tissue morphology, growth, and its selected endocrine functions.

1. Depots and cell types

Adipose tissue is divided into two distinct depots, subcutaneous and visceral. Subcutaneous adipose tissue, found directly below the skin, is an especially important heat insulator in the body, conducting only one third as much heat as other tissues. Visceral fat is located within the body cavity and surrounds many of the internal organs.

The distribution of fat in rodents has been thoroughly investigated, and was recently well described by Cinti et al. (5). Rodents have two main subcutaneous fat depots, one located anterior and one posterior. The anterior depot sits at the base of the forelimbs and primarily occupies the dorsal body region. Situated at the base of the hind legs, the posterior subcutaneous depot consists of a single tissue band beginning near the lumbar level. The visceral depots are located in the thorax and abdominal cavity. Within the thorax, visceral fat is located primarily among the intercostal nerve vascular bundles, the heart, and the aorta. Abdominal fat depots are either intra- or retroperitoneal. The retroperitoneal depot has an elongated conical shape and lies between the spine and the dorsal abdominal wall, while the perirenal depot lies next to the kidney. Perigonadal fat surrounds the ovaries, uterus, and bladder in females, while in males it surrounds the epididymis. The mesenteric depot is outlined by two peritoneal leaflets holding the intestine against the posterior wall.

The aforementioned depots are composed of a mixture of two distinct types of adipocytes, white and brown, which differ in morphology and function. Although both cell types can exist within the same depot, they are generally identified as two different tissues: 1) white adipose tissue (WAT), which appears yellow or white and contains predominantly white adipocytes, and 2) brown adipose tissue (BAT), which appears brown and contains mostly brown adipocytes.

The diameter of white adipocytes is variable, ranging between 30 and 70 μm according to depot site (6). Lipids within these cells are primarily organized within a large, unilocular droplet, the size of which can exceed 50 μm. Since their primary function is storage, white adipocytes are

12 spherical shape, allowing for maximal volume within a minimal space. The lipid droplet occupies the majority of intracellular space, forcing the cytoplasm and nucleus into a fine rim on the outer edge of the cell. Approximately 60 to 85% of the weight of WAT is lipid, with 90-99% being triglyceride. Small amounts of free fatty acids, diglyceride (DG), cholesterol, phospholipids and minute quantities of cholesterol ester and monoglyceride (MG) are also present. Conversely, brown adipocytes are polygonal, have centrally placed nuclei, and are relatively smaller than white adipocytes, ranging from 20 to 40 μm. These cells contain many large mitochondria packed with cristae within the cytoplasm, and lipids are stored in numerous small multilocular droplets (7). Although mature adipocytes constitute the main volume of adipose tissue, they are outnumbered by other cells present in this tissue, including fibroblasts, adipocyte precursor cells, and endothelial cells of the capillaries (5;7).

Although both adipocytes are capable of storing lipids in cytoplasmic droplets, their functions are different. WAT acts as both an endogenous energy store and an endocrine organ that secretes hormones that regulate appetite and energy homeostasis. The primary function of BAT is thermogenesis. Brown adipocytes uniquely express uncoupling protein-1 (UCP-1), which uncouples oxidative phosphorylation and utilize substrates to generate heat within the mitochondria (8). In this process, brown adipocytes hydrolyze stored triglycerides, releasing fatty acids that are immediately oxidized in mitochondria. The uncoupling protein blocks development of a H+ electrochemical gradient, thereby stimulating respiration. The free energy change associated with respiration is dissipated as heat.

Adipose tissue development begins in utero, with the adipocyte lineage derived from stem cell precursors that have the potential to become either brown or white adipose tissue (9). The details of this conversion into adipocytes (adipogenesis) will be discussed later in this section. Both WAT and BAT have critical functions at different stages of life. Fetal adipose tissue has the morphological and biochemical properties of brown adipose tissue, and is geared towards maximizing the abundance of UCP-1 (10). This is of particular importance in neonates, small mammals in cold environments, and in animals that hibernate (11). In humans and sheep, the abundance of brown fat is maximal around birth and is located primarily around core organs, enabling maximal heat production following cold exposure in the extrauterine environment. In these species, BAT is deteriorated after the postnatal period (12;13). Rodents maintain their body temperature by huddling together in a nest, and maximal UCP-1 expression occurs postnatally and is maintained throughout life (14). The chronology of WAT appearance is species-specific, and depends on the depot. WAT is not detected during embryonic life or at birth in rats or mice, but is present at birth in many species, including humans (15;16). After birth, WAT expansion takes place rapidly as a result of increased cell size and number.

Hormones such as glucocorticoids, triiodothyronine, epinephrine, and insulin-like growth factors, together with the developing sympathetic nervous system are essential for promoting prenatal adipose tissue deposition and ensuring maximal UCP1 abundance at birth (17). Among the factors that influence brown adipocytes, norepinephrine is best studied, revealing its significant physiological functions, not only for the acute thermogenic process but also for the control of cell proliferation, differentiation, and apoptosis (11).

13 All adipose tissue depots receive vascular and nerve supply. Collections of white adipocytes comprise fat lobules, each of which is supplied by an arteriole and surrounded by connective tissue septae. Blood is supplied to individual adipocytes by adjacent capillaries. Compared with WAT, BAT is more vascularized, being populated with multiple capillaries. This feature, combined with the high number of densely packed mitochondria in brown adipocytes results in its brown color. Both WAT and BAT are equally dependent on the blood outflow for delivery of adipocyte products to the organism. Through the use of anterograde and retrograde tracing techniques, direct SNS postganglionic innervation of WAT by noradrenergic fibers is now well established (18). BAT also makes synaptic contacts, and in contrast to WAT, is highly innervated (19). Both types of adipose tissue express α- and β-adrenergic receptors, with most adrenergic input to adipose tissue coming from the adrenal glands. (20)

2. Adipogenesis

In contrast to all other organs or tissues, adipose tissue is capable of seemingly unlimited growth in postnatal life. WAT is particularly unique in its potential for enormous changes in volume. With fat mass increasing up to 4-fold in morbidly obese humans, it can reach 60% to 70% of total body weight (21). WAT expansion results from both increased adipocyte size and number. Adipocyte hyperplasia has been observed in various rodent models, including rats fed a high-fat or high-carbohydrate diet, and transgenic mice overexpressing GLUT4 in adipose tissue (22;23). Although fat cell precursors from adult WAT can differentiate into mature adipocytes in vitro, adipocyte hyperplasia in humans remains controversial (24).

The molecular events leading to the commitment of embryonic stem cell precursors to the adipocyte lineage remain to be characterized. Studies using multipotent clonal cell lines indicate embryonic stem cell precursors have the capacity to differentiate into the mesodermal types of adipocytes, chondrocytes, osteoblasts, and myocytes (25). On the other hand, preadipocyte cell lines and primary adipocytes are already committed to the adipocyte lineage, thus representing different stages of adipocyte development.

The most commonly used preadipocyte cell lines are 3T3-F442A and 3T3-L1. These cells have been clonally isolated from disaggregrated 17- to 19- day Swiss 3T3 mouse embryos (26). In vitro-differentiated adipocytes have many characteristics of adipose cells in vivo. Subcutaneous injection of preadipocytes in nude mice leads to development of fat cell lobules that are histologically indistinguishable from WAT, suggesting that adipose cell acquisition occurs by a similar mechanism in vivo (27).

Preadipocyte differentiation is complicated, involving a precise chronology of morphological changes and gene expression. During the growth phase, preadipocyte cell lines as well as primary preadipocytes are morphologically similar to fibroblasts. They maintain the capacity for

14 Characteristics Cell Type Molecular events

Determined Preadipocyte Growth Arrest C/EBPβ Early Mitosis PPARγ Clonal expansion C/EBPα Late Committed Lipogenic enzymes

Terminal ECM alterations Differentiation & cytoskeletal remodeling

Secreted products Mature Adipocyte

Figure 1: Overview of adipocyte differentiation. Fibroblast-like preadipocytes are growth arrested by contact inhibition. When given appropriate adipogenic cues, preadipocytes undergo clonal expansion, followed by expression of early adipogenic genes, at which point the cells become committed to becoming adipocytes. The expression of lipogenic enzymes, storage of lipids, as well as extracellular matrix alterations, cytoskeletal remodeling and secretion of adipokines occur later, and are hallmarks of terminal differentiation.

15 growth, but must undergo growth arrest and withdraw from the cell cycle before conversion. Preadipocyte growth arrest is generally achieved through contact inhibition, but can be induced by maintenance in serum-free media (9). Following growth arrest the cells undergo at least one round of DNA replication and cell doubling, leading to clonal amplification of committed cells (28). After exiting the cell cycle, preadipocytes require a specific combination adipogenic cues to initiate differentiation. Although the full complement of inducing agents required for differentiation varies with each cell culture model, insulin, cAMP, and glucocorticoids are generally considered necessary for the induction of differentiation. However, the chronology of molecular events and pattern of gene expression can differ with culture models and differentiation protocols.

Terminal differentiation of adipocytes involves changes in the expression levels of ~300 genes/proteins. Several authors have attempted to schematize the steps of adipose conversion into a simple hierarchy of molecular events, with genes whose expression is modulated during adipogenesis categorized into early, intermediate and late mRNA and protein markers. However, adipocyte differentiation should be considered as a continuum of overlapping molecular events.

Expression of lipoprotein lipase (LPL) mRNA has often been cited as an early sign of adipocyte differentiation (25). LPL, discussed in detail later in this chapter, is secreted by mature adipocytes and plays a central role in controlling lipid accumulation. However, LPL expression occurs spontaneously at confluence without the addition of adipogenic agents, suggesting that LPL expression reflects the growth-arrest stage rather than an early differentiation step (29). Because LPL expression is not adipocyte-specific and is independent of the agents required for adipocyte differentiation, its classification as an early marker of adipocyte differentiation is questionable.

Several transcription factors, notably CCAAT-enhancer binding proteins (C/EBP-α, -β and -δ), and peroxisome proliferator-activated receptors (PPAR-γ and -β/-δ) are key regulators of adipogenesis (9). The PPARs belong to type II nuclear hormone receptor family and form heterodimers with the retinoid x receptor (RXR) (30). They are activated by a variety of structurally dissimilar compounds, notably the thiazolidones (TZD), and regulate transcription through binding of PPAR-RXR heterodimers to a response element. Members of the C/EBP family have a basic transcriptional activation domain and an adjoining leucine zipper motif, which provides the ability for homo- and heterodimerization.

C/EBP-β and -δ are first induced in response to an adipogenic cocktail, and in turn activate PPAR-γ2 and C/EBP-α expression (31;32). The importance of C/EBP-β and -δ during adipogenesis has been demonstrated using transgenic mice, with overexpression of either C/EBP-β or -δ in preadipocytes enhancing adipogenesis, whereas embryonic fibroblast cells derived from mice lacking either C/EBP-β or -δ do not differentiate efficiently (33). Mice lacking both C/EBP-β and -δ exhibit a decrease in WAT development, while embryonic fibroblast cells derived from these mice do not differentiate into mature adipocytes (33). C/EBP- β and -δ induction is followed by an increase in PPAR-γ expression. In mice, there are three PPAR-γ isoforms, PPAR-γ1, -γ2 and -γ3, which are generated from the same gene by alternative promoter usage and mRNA splicing (34). PPAR-γ2 is preferentially observed in adipocytes required for adipogenesis (35). Ren et al. demonstrated that PPAR-γ2-deficient pre-adipocytes,

16 fail to undergo differentiation while exogenous delivery of PPAR-γ2 into PPAR-γ-deficient cells completely restores adipogenesis.

C/EBP-α expression rises after PPAR-γ2 induction, just before the expression of adipocyte- specific genes. The requirement of PPAR-γ and C/EBP-α in adipose tissue development has also been demonstrated using transgenic knockout mice. Deletion of either gene results in a failure to develop normal adipose tissue (36;37). These two factors appear to co-regulate each other’s expression, as PPAR-γ heterozygous gene knockout mice display a sharp decrease in C/EBP-α levels, while PPAR-γ2 expression is reduced in C/EBP-α null animals (38). Interestingly, C/EBP-α-null embryonic fibroblast cells fail to undergo adipogenesis, an effect that can be restored by overexpression of PPAR-γ2. On the other hand, forced expression of C/EBP-α in PPAR-γ-null embryonic fibroblast cells does not allow the cells to differentiate (39). These experiments demonstrate that PPAR-γ2 is the key transcriptional regulator of adipogenesis, and that C/EBP-α may have an accessory function for maintaining PPAR-γ2 expression. Yet, PPAR- γ and C/EBP-α are essential transcription factors of adipogenesis that act in concert to generate fully mature adipocytes.

In addition to transcription factors, other signaling molecules regulate adipogenesis. For example, Pref-1, a plasma membrane protein containing six EGF repeats in the extracellular domain, is an inhibitor of adipocyte differentiation. Pref-1 is highly expressed in 3T3-L1 preadipocytes, but is not detectable in mature fat cells. Down-regulation of pref-1, which is induced by dexamethasone, is required for differentiation (40).

Induction of adipogenesis leads to major changes in cell shape and size. Differentiating preadipocytes transform from a fibroblast-like to a spherical shape, with dramatic changes occurring in cell morphology, cytoskeletal components, and the levels and types of extracellular matrix (ECM) components. These changes in cell shape reflect a distinct process in differentiation and are not the result of accumulated lipid stores. 3T3-L1 preadipocytes can undergo biochemical and morphological differentiation even under conditions whereby TG accumulation is blocked (41).

During the terminal phase of differentiation, adipocytes markedly increase de novo lipogenesis and acquire sensitivity to insulin. They form lipid droplets, which later fuse, resulting in the rounded, signet-ring appearance similar to those in WAT. There is a dramatic increase in the mRNA, protein, and activity of enzymes involved in TG metabolism, including ATP citrate lyase, malic enzyme, acetyl-CoA carboxylase (ACC), stearoyl-CoA desaturase (SCD1), glycerol- 3-phosphate acyltransferase, glycerol-3-phosphate dehydrogenase, fatty acid synthase (FAS), and glyceraldehyde-3-phosphate dehydrogenase (9). Furthermore, glucose transporters, insulin receptor number, and insulin sensitivity increase (42). During adipogenesis, there is also a loss of β1-adrenergic receptors and an increase in the β2- and the β3-subtypes, resulting in an increased number of total adrenergic receptors (43).

In addition to increases in proteins directly related to lipid and glucose metabolism, adipocytes also synthesize other adipose tissue-specific products, such as aP2, an adipocyte-specific fatty acid binding protein, and perilipin, a lipid droplet-associated protein (44). Furthermore, adipocytes produce a number of secreted products, some of which will be discussed later in this

17 chapter. These include adiponectin, resistin, fasting-induced adipocyte factor (FIAF). Leptin is also increased during in vitro terminal differentiation of adipocytes, although its level is much lower than that detected in adipose tissue (45). PPAR-γ and/or C/EBP-α are implicated in the activation of several of these genes, including aP2, GLUT4, and leptin (46).

The highly complex genetic reprogramming that occurs upon adipogenesis is under the tight control of hormones, cytokines, nutrients and signaling molecules that change the expression and/or activity of transcription factors that in turn finely regulate adipogenesis. The number of factors and their roles in this process are too numerous too mention here, and the reader is referred to the reviews by Gregoire and Harp (9;47). Briefly, GH, insulin, IGF-1, glucocorticoids, and agents that increase cAMP levels are generally considered positive effectors, while members of the TGF-β family, interferon-γ, interleukin-1β, and PKC inhibitors are viewed as negative regulators.

In conclusion, preadipocyte differentiation is a complex process, requiring up- and down- regulation of adipogenic and anti-adipogenic factors, respectively. Identification of the exact interactions between environmental signals, intracellular signaling pathways, and pro- or anti- adipogenic transcription factors remains a major issue. Furthermore, little is known about the signaling processes subsequent to cell-cell or cell-matrix interactions that modulate transcriptional activity and adipocyte differentiation. Understanding the various molecules that regulate adiposity and energy balance remains an important area for future research.

3. Lipid metabolism

Lipids are central building blocks of life. They are constituents of biological membranes, energy storage compounds, and messenger substances. As a source of energy, fat is the most concentrated, containing more than twice as much as proteins or carbohydrates. Thus, storing calories in the form of fat is an extremely efficient strategy, whether it is accomplished via FFA uptake from the blood, or by de novo synthesis of fatty acids (lipogenesis). In fact, the ability to store energy as fat probably offered distinct evolutionary advantages to animals.

While the brain requires glucose as a source of energy, many other tissues can utilize fat in the form of FFA as a source of energy. A portion of these FFA are obviously derived from ingested food; however, fat stores, mobilized by lipolysis, serve as an important source of energy between meals. This section will introduce the major factors that involved in lipid influx (lipoprotein lipase), lipogenesis (fatty acid synthase) and lipolysis (hormone sensitive lipase and perilipin).

Lipogenesis

Lipoprotein lipase (LPL) is a central enzyme in the metabolism of all classes of lipoproteins (48). Its initial discovery was in 1943 when Hahn et al. noted that injection of heparin into dogs led to a decrease in postprandial lipemia. Subsequent reports by Brunzell et al. described a “common, saturable, triglyceride removal mechanism” for chylomicrons and very low density lipoproteins (49).

18 LPL is now identified as the primary enzyme responsible for the hydrolysis of triglycerides to glycerol and FFAs for transport into metabolic tissues. LPL is synthesized by parenchymal cells, secreted as an active dimer, and is transferred to the luminal surface of endothelial cells where it binds to heparin sulfate proteoglycans. There, LPL is exposed to numerous circulating factors, including large multi-molecular lipoprotein particles, composed of TG, cholesterol, and apolipoprotein components that act as binding and enzyme activation sites (48). Endogenously produced triglycerides are secreted by the liver as very low-density lipoproteins (VLDL), while dietary triglycerides are incorporated into chylomicrons in the intestines (50;51). TG-rich lipoprotein particles, which are too large to cross the capillary endothelium in most tissues, bind to LPL on the capillary wall where they are enzymatically processed. Lipoprotein-bound LPL catalyzes the rate limiting step in the hydrolysis of TG within lipoproteins, producing 2- monoacylglycerol and non-esterified fatty acids (NEFA) for tissue utilization (Fig. 2).

In addition to its lipolytic activity, LPL possesses several physiological activities which all affect the plasma lipoprotein profile and the cellular metabolism of fatty acids and lipids. First, LPL is able to interact with lipoproteins independently of its lipolytic activity, by anchoring lipoproteins to the vessel wall and facilitating TG hydrolysis and lipoprotein particle uptake (52). Second, LPL acts as a ligand for the LDL receptor and several other members of the LDL receptor family, resulting in enhanced binding and uptake of lipoproteins (53). Lastly, LPL can mediate the selective uptake of lipoprotein associated lipids and lipophilic vitamins, without taking in lipoprotein particles (54).

The major sites of LPL production are adipose tissue, cardiac muscle, and skeletal muscle; however, LPL is expressed in other sites, including the nervous system, heart, adrenals, macrophages, proximal tubules of the kidneys, pancreatic islet cells, and lungs (55). Whereas LPL is absent from the liver of adult animals, hepatic LPL mRNA and enzyme activities are detectable during the suckling period and can be induced by cytokines such as TNFα (56). Tissue specific regulation of LPL expression provides a mechanism for localized control of the uptake of FFA, lipids and lipoproteins resulting in an effective distribution of nutrients and lipids among tissues. Perturbation of tissue-specific LPL expression has major metabolic consequences for energy homeostasis and lipoprotein metabolism. Mice lacking cardiac LPL have significantly elevated plasma TG levels and decreased clearance of postprandial lipids, while those completely lacking LPL develop severe hypertriglyceridaemia and die within twenty hours of birth (57). Dysregulation of LPL activity may be correlated with excessive lipid accumulation and has been proposed to play a role in the pathogenesis of obesity, atherosclerosis, and the development of insulin resistance (58).

The LPL gene spans ~30 kb and is divided into 10 exons, with substantial sequence homology among most of the species that have been examined (59). The cDNA codes for a 475-amino-acid protein including a 27-amino-acid signal peptide. The catalytic center is formed by three amino acids: Ser132, Asp156, and His241. Almost one hundred naturally

19 Insulin Catecholamines Glucagon ß-receptor Insulin PI3K Perilipin + PKA Adenosine Malonyl CoA + LPL LPL + FAS P P VLDL FFA HSL FFA FFA Triglycerides Glycerol Lipid droplet G3P Glycerol Glycerol Glucose

Endothelium

Figure 2: Schematic summarizing lipid flux in adipocytes. Triglyceride (TG) containing very-low density lipoproteins (VLDL) bind to lipoprotein lipase (LPL) inside the endothelium, where TG are cleaved into glycerol and free fatty acids (FFA) for entry into the adipocyte. LPL activity is decreased by glucagon and increased by insulin via PI3K. Once inside the adipocyte, FFA are either utilized for energy or combined with glycerol-3-phosphate (G3P) to reform TG, which enter the lipid droplet. During lipolysis, catecholamines bind to β-adrenergic receptors and stimulate PKA-mediated phosphorylation (P) of hormone sensitive lipase (HSL) and perilipin. At the lipid droplet surface, HSL catalyzes the cleavage of TG into FFA and glycerol. This process is inhibited by insulin and adenosine.

20 occurring mutations in the LPL gene have been described in humans, many of which are discussed in the review by Merkel et al. (60).

LPL activity is mainly regulated at both the transcriptional and translational levels (61). Numerous positive and negative regulatory sequences have been identified in the LPL promoter, which spans ~1.7 kb upstream of the transcription start site. The LPL promoter contains an octamer sequence motif (ATTTGGAT) which is sufficient for basal promoter function (62). Many other elements were discovered while studying 3T3-L1 differentiation, during which transcription of the LPL gene increases dramatically (61). Sites responsible for increases in LPL gene transcription during adipogenesis include sterol regulatory element binding protein 1-c (SREBP) sites and PPRE sites (63;64). GAS-like elements, a CCAAT box, and an inhibitory AP- 1-like sequence have recently been identified in the promoter region (65-67). LPL activity is also regulated at the translational level. Catecholamines and other inhibitors of LPL translation induce PKA-mediated production of an RNA-binding protein that binds to a region on the proximal 3'-untranslated region of the LPL mRNA (68).

Fatty Acid Synthase (FAS)

FAS is the key enzyme for de novo lipogenesis, catalyzing the reactions for the synthesis of long- chain fatty acids. This process involves a conserved set of chemical reactions for the cyclic stepwise elongation of activated precursors by two-carbon units. Although all organisms use variations of this common synthetic scheme, three distinct architectures for fatty acid synthesis have evolved. In bacteria, all reactions are carried out by individual, monofunctional proteins in a dissociated or type II FAS system. In contrast, the eukaryotic type I FAS consists of large, multifunctional polypeptides. Fungal FAS is a dodecamer that assumes a barrel-shaped structure, in which the catalytic domains are distributed over two distinct subunits (69). The FAS of animals is a homodimer of two 270-kD polypeptides that assembles in an X-shaped structure (70). It harbors all catalytic activities required for the synthetic cycle, making it one of the most complex mammalian enzymes.

Each subunit of mammalian and vertebrate FAS contains three N-terminal domains, the β- ketoacyl synthase, malonyl/acetyl transferase and dehydrase separated by a structural core from four C-terminal domains, the enoyl reductase, β-ketoacyl reductase, acyl carrier protein (ACP) and thioesterase. The thioesterase domain represents the enzyme activity required for releasing the mature fatty acid from the enzyme, while ACP is the site at which the growing fatty acid is bound during synthesis.

Fatty acid synthesis occurs in the cytosol via an anabolic pathway consuming 14 ATP and 7 NADPH per fatty acid synthesized. The key feature of the pathway for de novo biosynthesis of fatty acids is the sequential extension of an alkanoic chain, two carbons at a time, by a series of decarboxylative condensation reactions that can be summarized by the equation:

+ Acetyl-CoA+7 Malonyl-CoA+14 NADPH+14 H → Palmitic acid +7 CO2+8 CoA+14 + NADP +6 H2O

21 Under conditions that favor fatty acid synthesis, mitochondrial citrate synthase catalyzes the formation of citrate from acetyl-CoA and oxaloacetate. Citrate exits the mitochondria via the tricarboxylate anion carrier and is cleaved to acetyl-CoA by ATP citrate lyase. Acetyl-CoA, the precursor for fatty acid synthesis, is converted to malonyl-CoA by ACC, which is the rate limiting enzyme in the fatty acid synthesis pathway. ACC activity is regulated by substrate supply, allosteric activation by citrate and inhibition by fatty acyl-CoA, and phosphorylation of multiple serine residues and interactions with other proteins.

The reaction cycle of FAS is initiated by the transfer of the acyl moiety of acetyl-CoA to the ACP. ß-ketoacyl-ACP, an intermediate produce, is processed by nicotinamide adenine dinucleotide phosphate (NADPH)–dependent reduction. The resulting ß-hydroxyacyl-ACP is dehydrated and then reduced by the NADPH-dependent ß-enoyl reductase to yield a four-carbon acyl substrate for further cyclic elongation with two-carbon units until a substrate length of C14 (myristate), C16 (palmitate) or C18 (stearate) is reached. Finally, the product is released from the ACP by the thioesterase.

In humans, FAS expression is ubiquitous; with the highest levels in liver and lung. FAS mRNA levels are high in lipogenic rodent tissues, liver and adipose tissue (71). The mammalian FAS gene is responsive to nutritional and hormonal cues. FAS is undetectable in the livers of fasted mice and is dramatically induced upon refeeding a high carbohydrate, fat-free diet (72). Nutritional regulation of FAS occurs mainly via changes in FAS gene transcription. Insulin secretion, which is increased during feeding, has a positive effect on FAS expression (73). Glucagon, acting through cAMP-dependent mechanisms, suppresses FAS, while thyroid hormone increases expression (74;75).

The FAS promoter contains numerous binding sites for stimulatory proteins (Sp1/Sp3) as well as cis-elements corresponding to nuclear factor Y (NF-Y), upstream stimulatory factor (USF) and SREBP (76). Multiple insulin response elements have been identified in the promoter region, while a putative carbohydrate response element (CRE) site is located in intron 1 (76). USF and SREBP-1c seem to play an important and possibly cooperative role in regulating FAS transcription. Moon et al. have demonstrated that binding of SREBP to the sterol response element (SRE) is responsible for low-level induction, whereas occupancy of an E-box by USF is necessary for high-level activation of the FAS promoter (77). Furthermore, FAS induction by high carbohydrate/insulin is severely impaired in USF and SREBP knockout mice, demonstrating a requirement for both of these transcription factors (78).

Hormone Sensitive Lipase (HSL)

The primary sources of fatty acids for oxidation are dietary and mobilization from cellular stores. Fatty acids from the diet are delivered from the gut to cells via transport in the blood. Fatty acids are stored in the form of TG primarily within adipocytes of adipose tissue. In response to energy demands, the fatty acids of stored TG can be mobilized for use by peripheral tissues.

HSL is the enzyme that catalyzes intracellular TG hydrolysis, and is a major regulator of fatty acid mobilization in adipose tissue as well as other tissues. The first reports of hormone- responsive lipolytic activity in adipose tissue occurred in the early 1960s, when a lipolytic

22 activity with properties different from those of LPL was described (79). It soon became evident that hydrolysis of TGs occurs through three consecutive reactions and is catalyzed by two enzymes: HSL and monoglyceride lipase (MGL). HSL alone catalyzes the hydrolysis of TGs and diglycerides (DG), whereas the participation of an additional enzyme, MGL is required to obtain complete hydrolysis of monoglycerides (MG) (80).

The gene structure of human and mouse HSL are very similar (81). Both genes comprise nine main coding exons, spanning approximately 11 kb. In humans, four additional exons, designated T1, T2, A, and B, lie approximately 16, 12, 12.5, and ~1.5 kb upstream of exon 1, respectively, and differentially code for 5’-UTRs (82). Only the smallest HSL mRNA product, transcribed from exon B, is expressed in human adipose tissue, while exons T1 and T2 are transcribed in the testes (83). In mouse HSL, five alternative noncoding exons (A, B, C, D, and 1) have been reported upstream of the translation start site, each of which can be alternatively utilized and expressed in mouse adipose tissue to varying degrees (84).

The HSL protein exists as several isoforms that are differentially expressed in a number of tissues. The major isoform, produced in adipose tissue, is a single polypeptide with a molecular mass of 84- and 88 kDa in rodents and humans, respectively. At least three additional isoforms have been reported. The testes express two isoforms, an 84 kDa protein similar to adipose HSL, and a larger ~120-130 kDa product. Pancreatic islets and ß cells have an HSL isoform that contains an additional 43 amino acids N-terminal to the normal adipose form (85).

Structurally, HSL contains two major domains: a C-terminal catalytic domain that contains a regulatory module and an N-terminal domain involved in protein-protein and protein-lipid interactions (86). The N-terminal region shows high conservation (91%) between man and rodents, but is variable between isoforms (87). This is followed by a short stretch of poorly conserved amino acids believed to be a linker between domains. The C-terminal catalytic domain is identical in all known HSL isoforms. It contains the active site, which is a catalytic triad of serine, aspartic or glutamic acid, and histidine, as well as a regulatory region that contains all known phosphorylation sites on the protein. HSL exists as a functional dimer composed of homologous subunits, and has significantly greater hydrolytic activity but no difference in substrate affinity when compared with monomeric HSL (88).

Compared with other lipases, HSL has broad substrate specificity. It hydrolyzes TGs and DGs, as well as cholesteryl esters, steroid fatty acid esters, retinyl esters, and para-nitrophenyl esters (89). However, in contrast to many other lipases, HSL has no phospholipase activity. The lipolytic activity of HSL against DGs is higher than that against TGs by 10-fold. This, in combination with the action of MGL, assures no accumulation of intermediate metabolites (90). Lipolytic activity against cholesteryl esters is about twice the activity toward TG, while the esterase activity is more than 20-fold that of TGs (87). HSL has high specificity for the 1- or 3- ester bond, while MGL is believed to be mainly responsible for hydrolysis of the ester bond in the 2- position (91).

HSL activity is primarily activated by PKA-mediated phosphorylation of serine residues in the regulatory region. In rodents, at least three serine residues, S563, S659, and S660 have been implicated in PKA-mediated function (92). Lipolysis is acutely regulated by hormones,

23 neurotransmitters and other effector molecules. Catecholamines are important stimulators of lipolysis, whereas insulin is the most effective anti-lipolytic hormone. Binding of catecholamines to the β-adrenergic receptors, coupled to adenylate cyclase via the stimulatory G-protein, leads to increased production of cAMP and activation of PKA. Lipolytic hormones also activate the MAPK pathway, which regulates lipolysis by phosphorylating S600 in the regulatory region of HSL (93). Other kinases such as glycogen synthase kinase-4, Ca++/calmodulin-dependent protein kinase II, and AMPK phosphorylate HSL at a basal activation site, S565 in rat HSL (94).

In addition to the activation of HSL hydrolytic activity, other mechanisms involving HSL have been suggested to account for lipolysis. Several studies indicate that catecholamine-induced stimulation of lipolysis in 3T3-L1 adipocytes or in rat adipocytes is due to the translocation of phosphorylated HSL from a cytosolic compartment to the lipid droplet (95). However, this does not appear to be true for all physiological conditions, since translocation of HSL was not observed with lipolytic stimulation in adipocytes from old or lactating rats (95).

HSL mRNA is expressed in a variety of tissues, including white and brown adipose tissue, steroidogenic tissues, mammary gland, muscle tissues, macrophages, and endocrine pancreas. In rat, HSL expression is lower in subcutaneous fat compared with internal fat depots, suggesting a possible basis for the differences in the rate of lipolysis among various fat depots (96).

HSL-knockout models have clarified the physiological importance of this enzyme. Fortier et al. report that HSL-deficient mice have WAT abnormalities, including low mass, marked heterogeneity of cell diameter, increased DG content, and low ß-adrenergic stimulation of adipocyte lipolysis. WAT expression of normal HSL markedly improves HSL knockout WAT biochemically, physiologically, and morphologically (97). Male HSL -/- mice are sterile because of oligospermia. Their testes completely lack neutral cholesterol ester hydrolase activities and contain increased amounts of cholesterol ester (98).

Perilipin

The mechanism for catecholamine-induced lipolytic action was originally believed to result solely from phosphorylation and activation of HSL by PKA. However, this explanation was not in accord with observations made by Frederikson and Nilsson. In these studies, a modest 2-3 fold activation of HSL catalytic activity was observed in vitro, whereas a remarkable 50 to 100-fold induction of lipolysis was observed upon PKA activation in intact adipocytes (89). Subsequent studies of HSL localization demonstrated that the enzyme was found in the cytosol of unstimulated adipocytes, whereas upon lipolytic stimulation, the lipase became tightly associated with the lipid storage droplet (99). Immunofluorescence studies using anti-HSL antibodies in 3T3-L1 adipocytes confirmed these observations, and it was hypothesized that interactions between a catalytically active HSL and neutral lipid stores at the lipid droplet (LD) surface accounted for the increases in catecholamine-stimulated lipolysis (100). However, the movement of HSL was not found to be associated with any cytoskeletal component, and it was not until recently that the critical role of the protein, perilipin A, in both HSL translocation and lipolysis was determined (101).

24 The perilipins are a family of closely related proteins that are abundantly expressed in adipocytes, and are concentrated at the surface of lipid droplets. Perilipin exists as four isoforms, A, B, C, and D as the result of alternative splicing of a single gene (102). Perilipin A and B are the predominant isoforms of murine adipocytes, with perilipin A constituting ~85% of total perilipin. The lesser investigated perilipin C is expressed only in steroidogenic cells, while perilipin D is found in steroidogenic cells and adipocytes. Perilipin A and B share a common N- terminal region of 1–405 amino acids but possess unique C termini (103).

Besides HSL, perilipins are major targets for PKA phosphorylation upon β-adrenergic stimulation of adipocytes (104). A function for perilipin was first described in 1998, when Souza et al. reported that the lipolytic action of TNF-α occurs via decreases of perilipin A at the surface of the lipid droplet, an action that was prevented by retroviral overexpression of perilipin (105). These observations suggested that perilipins serve as suppressors of basal lipolysis. Further support for this function came from Brasaemle and Tansey, who noted that expression of perilipin A in Chinese hamster ovary cells inhibited TG hydrolysis by 87% when PKA was quiescent (100;106). Interestingly, they previously reported that adipocytes lacking perilipin A exhibited a near total loss of ß-adrenergic receptor-stimulated lipolysis, despite the presence of normal levels of HSL protein per cell (107). These data suggest that perilipin A is not only required to maintain adipose cells in the quiescent state, but is elicits a functional lipolytic activation in adipose cells. Indeed, Stzalyrd et al. have demonstrated that HSL fails to translocate to lipid droplets in adipocytes lacking perilipin A (101). Moreover, phosphorylation of perilipin A appears necessary for proper function, as mutation of PKA-mediated phosphorylation sites within the N-terminal region of perilipin A abrogates the PKA-mediated lipolytic response (106).

There is some recent controversy over the role of perilipin A in HSL transport. Miyoshi et al. report that PKA-stimulated HSL translocation is normal in adipocytes from perilipin A-deficient mice that express a mutant perilipin A lacking all six PKA phosphorylation sites (108). This data disagrees with the general belief that PKA-mediate phosphorylation of perilipin A is necessary for HSL translocation. This may be true, since this is the first study using perilipin A incapable of being phosphorylated.

Few studies have investigated perilipin B. Although perilipin B comprises <10% of total perilipin, it is the major isoform associated with the plasma membrane of human adipocytes (109). Aboulaich et al. recently reported that this association is controlled by insulin and catecholamines, as perilipin B was specifically depleted from the plasma membrane in response to isoproterenol, while insulin increased the amount of threonine phosphorylated perilipin at the plasma membrane (110). In addition, it was noted that perilipin at the plasma membrane was phosphorylated on a cluster of three threonine residues. The authors suggest that the hormone- controlled association of perilipin B with the plasma membrane may act to protect the newly formed TG in the plasma membrane from hydrolysis.

Despite the central role of HSL in adipocyte lipolysis, the available evidence suggests that HSL is not the only hormone-responsive lipase in adipocytes. Mice with a targeted deletion of HSL have reduced catecholamine-sensitive adipocyte lipolysis but are not obese (98). In addition, both adipose tissue and isolated adipocytes derived from mouse embryonic fibroblasts of HSL- deficient mice have significant residual TG lipase activity, suggesting the presence of one or

25 more additional TG-specific lipases (111). Furthermore, this TG lipase activity is sensitive to regulation by various factors including catecholamines, tumor necrosis factor-α, and thiazolidinediones, suggesting that activity is also hormone responsive (111).

Indeed, a novel adipocyte triglyceride lipase has been recently identified, designated as adipose triglyceride lipase (ATGL), desnutrin in mice and calcium-independent phospholipase A2

(iPLA2)ζ in humans (112). To date, there is limited information available, most of which comes from experiments in 3T3-L1 cells, on the function of this lipase. ATGL is expressed abundantly in adipose tissue, but is found at a very low, barely detectable level in liver, spleen, thymus, kidney, brain, skeletal muscle, and lung (113). It is expressed at low but detectable levels early in differentiation and begins to increase within the first 24 hours of differentiation. This early expression precedes the expression of several adipogenic genes, including C/EBP-α and PPAR-γ, suggesting a potential role for ATGL in the adipogenic process (114). ATGL expression appears to be nutritionally regulated, as mRNA levels increased by fasting and decreased by re-feeding (113;114).

Current data suggests that HSL and ATGL may work in a serial rather than in a parallel fashion, with ATGL-mediated TG hydrolysis providing DG substrate for HSL, thus driving subsequent steps in the lipolytic pathway. This conclusion is supported by studies which show that ATGL increases hydrolysis of TG but not of other substrates, whereas HSL has broad substrate specificity and a 10-fold greater activity against DG than TG (112).

Villena et al. demonstrated that dexamethasone, but not the cAMP-mediated hormone glucagon, induced expression of ATGL mRNA in 3T3-L1 cells in a time- and dose-dependent manner in the KD range for the glucocorticoid receptor (113). Thus, it has been proposed that during fasting, plasma glucocorticoid levels increase, leading to the rise of ATGL expression in adipose tissue (115). Conversely, treatment of 3T3-L1 adipocytes with insulin, which favors energy storage, produces a dose-dependent decrease in ATGL expression (114). Taken together, the available data suggest that ATGL participates in TG-specific hydrolysis and hence, like HSL, may play an important role in lipid metabolism. However, although HSL acutely affects lipolysis, primarily via posttranslational mechanisms, ATGL may mediate more long-term effects on lipolysis via transcriptional mechanisms.

4. Glucose Transport

Following a meal, insulin secretion from the pancreas facilitates the removal of glucose from the bloodstream. One of the main events in this regulatory process is the stimulation of glucose entry into muscle and fat cells. This process is mediated by facilitative glucose transporters (GLUTs), which belong to an extended family of hexose transporters has evolved to meet the specific metabolic needs of various tissues (116). There are some fourteen members of the facilitative glucose transporter gene family that are divided into three classes based on conserved structural characteristics (117). Class I includes GLUTs 1–4, the best characterized transporters of the family, while Class II is comprised of GLUT5 (a fructose-specific transporter), and GLUTs 7, 9, and 11, which remain poorly understood. Class III includes GLUTs 8, 10, 12, and the proton- myoinositol symporter H+-myo-inositol cotransporter, all of which are still largely

26 uncharacterized. White adipocytes appear to express as many as eight members: GLUT1, GLUT3, GLUT4, GLUT5, GLUT8, GLUT10, and GLUT12 (118).

GLUT proteins transport glucose down its concentration gradient in an energy-independent manner. As most mammalian cell types are net consumers of glucose, they maintain low intracellular glucose concentrations, thus favoring glucose entry. One GLUT isoform in particular, GLUT4, is insulin responsive, acting as the primary regulator of glucose absorption in muscle and adipose. GLUT4 has been localized to tubulovesicular structures in the perinuclear region and distinct foci scattered throughout the cytosol (119). Perinuclear GLUT4 partially colocalizes with markers of the endosomal recycling compartment (ERC), the Golgi complex, and the trans-Golgi network (120). Using static microscopy analysis, it is not possible to discern whether perinuclear or cytosolic depots encompass the insulin-sensitive GLUT4-donor compartment. As a result, there is an ongoing controversy regarding the mechanism of GLUT4 trafficking/retention. One model proposes that GLUT4 retention is dynamic, based on slow exocytosis and rapid internalization of the entire pool of GLUT4 (121). In this model, insulin increases GLUT4 in the plasma membrane by modulating GLUT4 exocytosis and endocytosis. The second model is that GLUT4 retention is static, with ~90% of GLUT4 stored in compartments that are not in equilibrium with the cell surface in basal conditions (122). In this model, insulin increases GLUT4 in the plasma membrane by releasing it from a highly- responsive GLUT4 storage vesicle (GSV) compartment. Watson and Pessin agree with the dynamic model, based on observations that GLUT4 compartments can be co- immunoprecipitated with constitutively recycling proteins (123). Furthermore, recent measurements of GLUT4 trafficking kinetics in 3T3-L1 adipocytes with green-fluorescent protein fused to the carboxyl terminus found that GLUT4 retention is maintained by a dynamic mechanism involving slow exocytosis and rapid internalization (124). In support of the static model is the presence of insulin-responsive vesicular cargo compartment during early 3T3-L1 differentiation (125).

It is now widely accepted that activation of the insulin receptor tyrosine kinase leads to the translocation of the facilitative GLUT4 from intracellular stores, to the plasma membrane in adipose and muscle tissue, and that this effect contributes to the postprandial maintenance of the normal glycemic state. This redistribution corresponds to a shift from less than 5% GLUT4 in the plasma membrane in basal conditions to about 50% of GLUT4 in the plasma membrane in the presence of insulin. To understand how insulin leads to the translocation of GLUT4 to the cell surface of insulin-sensitive cells, it is necessary to identify a link between the insulin-signaling pathways and GLUT4 transport machinery. This has proven challenging, as although the signaling molecules that function proximal to the activated insulin receptor have been well characterized, it is not known how the distal insulin-signaling cascade interfaces with and mobilizes GLUT4-containing compartments.

Molecularly, GLUT4 translocation occurs through a signal-transduction cascade that converts the initial tyrosine kinase signal into a phosphoinositide intermediate that, in turn, activates a serine/threonine kinase cascade (Fig. 3). The insulin receptor (IR) is a heterotetrameric bifunctional complex, consisting of two extracellular α subunits that bind insulin, and two transmembrane β subunits with tyrosine kinase activity. Binding of insulin to the α subunit induces the transphosphorylation of one β subunit by another on specific tyrosine residues in an

27 activation loop, resulting in the increased catalytic activity of the kinase. In addition, several cytosolic scaffold proteins serve as substrates for the activated insulin receptor, thereby greatly expanding the repertoire of potential downstream signaling opportunities.

With respect to glucose uptake, four members of the insulin receptor substrate family (IRS1 through 4), Cbl, and APS undergo tyrosine phosphorylation in response to insulin stimulation (126). Insulin-induced tyrosine phosphorylation of IRS proteins generates docking sites for many SH2-domain-containing downstream effectors, particularly type-1A PI3K. Type-1A PI3Ks are heterodimers composed of a regulatory p85 subunit and a catalytic p110 subunit (127). The p85 subunit contains SH2 domains that bind phosphotyrosine residues in IRS and other proteins, thereby allosterically regulating the activity of the p110 catalytic subunit. Activated PI3K catalyses the conversion of phosphorylate phosphatidylinositol 4,5-bisphosphate (PIP2) to form phosphatidylinositol 3,4,5-trisphosphate (PIP3). This interaction of PI3K with IRS apparently provides a dual function by both activating the PI3K and targeting it to the plasma membrane localized substrate PIP2. PI3K-mediated increases of PIP3 in the membrane are thought to form lipid-based platforms needed for the recruitment and activation of a subset of signaling proteins with pleckstrin homology (PH) domains. These include protein serine-threonine kinases, protein tyrosine kinases (Tec family), exchange factors for GTP-binding proteins (Grp1 and Rac exchange factors), and adaptor proteins, such as GAB-1. PIP3 promotes the cell-surface localization of two PH-domain-containing enzymes involved in GLUT4 translocation, the 3- phosphoinositide-dependent protein kinase-1 (PDK1) and Akt. In a mechanism yet to be elucidated, these signaling cascades are thought to converge in a concerted manner to mobilize GLUT4-storage compartments (123).

Although PI3K appears to be necessary for insulin-stimulated GLUT4 translocation, substantial evidence indicates that the generation of PIP3 is not sufficient to recruit GLUT4 to the plasma membrane (128). Indeed, there is evidence that PI3K-independent mechanisms participate in the regulation of insulin-mediated GLUT4 translocation. Recent studies have shown that tyrosine phosphorylation of Cbl, resulted in the insulin-dependent recruitment of Cbl into caveolin- enriched lipid raft compartments (129). This process involves two adapter proteins, APS and cb1-associated protein (CAP) (130). CAP/Cbl heterodimerize and are recruited to activated IRs through interactions with APS (131). APS is also an IRS, and tyrosine-phosphorylated APS facilitates the phosphorylation of Cbl by the IR. Once Cbl is phosphorylated, the CAP/Cbl heterodimer appears to dissociate from the IR and accumulate in lipid raft domains. Dominant interfering mutants of CAP that prevent the localization of Cbl to lipid rafts specifically block insulin-stimulated GLUT4 translocation and glucose uptake, without affecting the PI3K signaling pathway (129). The mechanism by which CAP/cbl induces GLUT4 translocation is not entirely understood, but may involve one or more recently identified signaling molecules, including AS160, PIKfyve, and/or Synip.

The importance of functional IR signaling has been demonstrated in various knockout models. IR-deficient mice develop early postnatal diabetes and die of ketoacidosis, while ablation of either IRS1 or IRS2 leads to peripheral insulin resistance (132). IRS3 deficiency results in no apparent abnormalities and does not appear to be functional in humans (133). IRS4 knockout mice showed mild growth defects and were slightly glucose intolerant (134).

28 Insulin receptor

CAP PIP2 PIP3 Cbl P PDK P P PI3K Akt P APS P P IRS-1 P

CAP AS160 Cbl PI3K PIKfyve Synip ? Glucose

P Phosphotyrosine

GLUT4 GLUT4 vesicle GLUT4 endosome Lipid raft

Figure 3: Schematic of insulin-stimulated glucose uptake. Binding of insulin to its receptor induces the phosphorylation of insulin-responsive substrate-1 (IRS-1) and adaptor molecule containing pleckstrin homology and SH2 domains (APS). IRS-1 activates phosphatidylinositol-3-kinase (PI3K), which stimulates the conversion of phosphorylate phosphatidylinositol 4,5-bisphosphate (PIP2) to phosphatidylinositol 3,4,5-trisphosphate (PIP3). The formation of PIP3 activates Akt and phosphoinositol-dependent kinase (PDK), which increase glucose transporter 4 (GLUT4) vesicle translocation to the plasma membrane via several candidate molecules (AS160, PIKfyve, and Synip). APS phosphorylates the Cbl- associated protein (CAP)/Cbl complex, which translocates to lipid rafts in the membrane and stimulates GLUT4 trafficking via an unknown mechanism.

29 2. Endocrine function

WAT was once considered an inert energy storage compartment. However, investigations of the regulation of food intake in surgically joined parabiotic animals, which allows circulating substances to move between animals, supported the hypothesis that a humorally transported factor(s) is involved in regulating feeding behavior. For example, when one member of a pair of parabiotic rodents is overfed via stomach tube, the other member of the pair reduces its voluntary food intake (135). The potential humoral signals identified by such experiments were postulated to be nutrients, nutrient metabolites, or hormones.

Pioneering work from the Spiegelman and Flier in 1987 demonstrated for the first time that adipocytes are an abundant source of a specific secretory protein, called adipsin (136). Adipose tissue is now recognized as a major secretory organ, producing a variety of bioactive peptides known as adipokines which act at both the local (autocrine/paracrine) and systemic (endocrine) level. To date, there have been in upwards of fifty adipokines identified, far too many to discuss here. Instead, we will introduce only those factors most relevant to this thesis.

Leptin

Leptin is a 16-kDa polypeptide that has structural homology to cytokines. It is probably the most well-known adipokine, and has been reported to exert profound effects on energy homeostasis, neuroendocrine, immune, and cardiovascular systems. The effects of leptin were first observed over fifty years ago in mice with random gene mutations, designated ob (obese) and db (diabetic), resulting in obesity and hyperphagia. The product of the ob gene was discovered through positional cloning of the ob gene in 1994 by Jeffrey Friedman’s laboratory (137).

Mouse and human ob genes have been localized to chromosomes 6 and 7q31.3, respectively (137). Sequence homology is high across species, with human leptin being 84% identical to mouse leptin and 83% to rat leptin. The ob gene consists of 3 exons separated by 2 introns, and the coding region for Ob protein is located in exons 2 and 3. Transcription of the ob gene yields a mRNA of ∼3.5 kb that is expressed primarily in adipose tissue, but recent studies have confirmed that some other tissues also express leptin, including placenta, ovaries, skeletal muscle, hypothalamus, pituitary and stomach. Several regulatory elements have been identified within the ob gene promoter, including cyclic AMP and glucocorticoid response elements, and CCATT/enhancer and SP-1 binding sites.

The product of the ob gene is a 167 amino acid protein with an N-terminal secretory signal sequence of 21 amino acids. The signal sequence is functional, and results in the trans-location of leptin into microsomes with the subsequent removal of the signal peptide (137). NMR analysis showed that it was four-helix bundle cytokine, with helix lengths and disulfide patterns in agreement those of the short-helix cytokine family (138). Notably, this protein is comparable in weight and length, and is organized much like PRL. Furthermore, the protein structure of PRL and leptin bear striking similarities.

The protein was subsequently named leptin (from the Greek root leptos meaning thin) because it caused marked reductions in food intake, body weight, and body fat when injected into ob/ob or

30 normal mice (139). Leptin did not, however, exert these effects in db/db mice, which were later shown to have a defect in the leptin receptor (140).

Leptin signals through the leptin receptor (Ob), a member of cytokine receptor family and product of the diabetes (db) gene (141). Much like its ligand displays similarities to PRL; the Ob-R shares several characteristics with the PRLR. In mice, six alternatively spliced isoforms (Ob-Ra,-Rb,-Rc,-Rd,-Re and -Rf) have been identified, all of which are products of the single ob-r gene (142). There are four known isoforms of the human leptin receptor with different C- terminal cytoplasmatic domains designated by the number of their unique c-terminal amino acids Ob-R5,-15, -67 and 274 (143). Similar to the PRLR, all Ob-R isoforms have identical extracellular ligand binding domains but differ in their intracellular C-terminus. Ob-R isoforms can be divided into three classes: long, short, and soluble isoforms. The long isoform (Ob-Rb) is considered to be the major signaling isoform since it is the only isoform that can transmit the leptin signal via the complete Jak/Stat3 pathway (144). Ob-Rb is expressed throughout the body and has been located in monocytes, lymphocytes, pancreatic beta cells, enterocytes, endothelial smooth muscle, and other cells (145). Leptin binding has been shown in lung, intestine, kidney, liver, skin, stomach, heart, spleen (146). The expression of Ob-Ra is ubiquitous, and its proposed role includes the transport of leptin across the blood–brain barrier. In mice, alternative splicing results in different Ob-R variants including its soluble isoform, which potentially modulates steady-state leptin levels by binding free leptin in the circulation, and consequently preventing the hormone from degradation and clearance (147).

Numerous studies have reported that circulating leptin concentrations are highly correlated with indices of adiposity, such as BMI, percentage body fat and total fat mass (148;149). Furthermore, adipocyte size is an important determinant of leptin synthesis, as larger adipocytes contain more leptin than smaller adipocytes in the same individual (150). Comparison of ob mRNA levels in different adipose tissue depots suggests that there are site-specific variations in leptin expression. In humans, leptin expression is greater in subcutaneous than in visceral adipose tissue (151). Leptin levels exhibit sexual dimorphism in humans, with its circulating levels significantly higher in women than men after correcting for total body fat (152). The gender difference is reversed in rats, with male rats having higher leptin concentrations than female rats, reflecting the greater amount of body fat in male rats (152;153).

Adipose tissue mass is not the only determinant of circulating leptin concentrations. Fluctuating energy intake also has a major influence on plasma leptin levels. Plasma leptin levels are markedly lowered by fasting or dieting and are recovered during refeeding (154). Notably, these changes in leptin levels are disproportionate to the relatively small changes in body fat induced by these short-term interventions, suggesting that leptin serves as an indicator of energy stores, as well as a mediator of energy balance (155).

Support for this notion comes from observations in ob/ob mice, which show many of the abnormalities seen in starved animals, including decreased body temperature, hyperphagia, decreased energy expenditure, decreased immune function, and infertility. Leptin replacement corrects all of these abnormalities, implying that ob/ob mice exist in a state of 'perceived starvation' and that the resulting biological response in the presence of food leads to obesity (156). Furthermore, the report that exogenous leptin attenuates the neuroendocrine responses to

31 food restriction, such as the delay in ovulation in female mice and changes in gonadal, adrenal and thyroid axes in male mice, agree with the idea that decreased plasma leptin levels signal nutrient deprivation (157). Notably, these responses are readily normalized by low-dose leptin replacement.

In contrast, common forms of obesity are characterized by elevated circulating leptin. Neither endogenously high leptin levels nor treatment with exogenous leptin is effective in ameliorating this obesity, consistent with a state of leptin resistance (158). The mechanism for leptin resistance is unknown, but might be the result of reduced brain leptin transport, receptor dysfunction, or blunting of leptin signaling in the hypothalamus and other CNS targets (159).

Clearly, the functions of leptin are not limited to energy homeostasis, since leptin deficiency in ob/ob mice is associated with activation of the hypothalamo-pituitary-adrenal (HPA) axis and suppression of the hypothalamo-pituitary-thyroid and -gonadal axes. Leptin treatment decreases hypercortisolemia in ob/ob mice, inhibits stress-induced secretion of hypothalamic corticotrophin-releasing hormone (CRH) in mice, and inhibits cortisol secretion from adrenocortical cells in vitro (160). The role of leptin in HPA activity in humans in vivo remains unclear. Leptin is also important in regulating the onset of puberty. Treatment of mice with leptin accelerates the maturation of the female reproductive tract and leads to an earlier onset of the estrous cycle and reproductive capacity (161).

Leptin regulates specific neuronal groups within the hypothalamus, brainstem and other regions of the CNS. Leptin receptors, located throughout the brain, are highly expressed in the arcuate, dorsomedial, ventromedial and ventral premamillary hypothalamic nuclei, and colocalize with neuropeptides involved in energy homeostasis (162). Notably, Ob-Rb is expressed in neurons within the medial arcuate nucleus that produce the orexigenic factors Neuropeptide Y (NPY) and agouti-related protein (AgRP). An increase in leptin directly suppresses the production of these peptides (163). Other orexigenic peptides, such as melanin-concentrating hormone (MCH) and orexins are synthesized in the LHA and are inhibited indirectly by leptin (164). Leptin increases the levels of anorectic peptides, including α-melanin-stimulating hormone (α-MSH), derived from proopiomelanocortin (POMC), and cocaine and amphetamine-regulated transcript (CART), in the arcuate nucleus, as well as CRH in the PVN (165).

Interestingly, Simerly et al. have recently suggested a developmental role for leptin in the brain, where it may direct the development of hypothalamic pathways involved in energy homeostasis by promoting axonal projections from the arcuate nucleus of the hypothalamus to other hypothalamic sites that mediate the effects of leptin on food intake and body weight (166).

In addition to its central effects, a growing body of evidence has indicates that leptin also directly acts on ß-cells. Leptin exerts a long-term control of insulin secretion, thereby adapting the amount of insulin secretion to the amount of body fat stores (167). However, this tonic restriction of insulin secretion by leptin does not interfere with the short-term stimulatory actions of nutrients and hormones, such as glucose- and incretin-dependent insulin secretion. Conversely, insulin stimulates leptin secretion from WAT following energy intake (168). This establishes a classic endocrine feedback loop—the so-called "adipo-insular axis.”

32 Some important differences between mice and humans deficient in leptin are now apparent. Leptin-deficient mice have clearly reduced energy expenditure and efficient metabolism such that obesity develops even without overeating. In contrast, efficient metabolism has not yet been documented in several totally leptin deficient humans; hyperphagia appears to be the sole or dominant factor (159).

Adiponectin

Adiponectin, (adipocyte complement-related protein) also known as ACRP30, GBP28 (gelatin binding protein 28), apM1 (adipose most abundant gene transcript 1), and AdipoQ, is an adipocyte-specific factor that was discovered in 1995/1996 by several independent groups (169- 172). Developmentally, murine adiponectin mRNA is first detected during late embryogenesis (day 17), in a pattern consistent with that seen in the mouse for other adipocyte markers during embryogenesis (173). Since its discovery, adiponectin has been investigated as a potential antidiabetic, anti-atherosclerotic, and anti-inflammatory agent.

In its monomeric form, adiponectin is approximately 30 kDa and in composed of 247 amino acids. It consists of four domains, an amino-terminal signal sequence, a variable region, a collagen-like domain that is important for building secondary and tertiary structure, and a C- terminal globular domain that is responsible for mediating its effects (169). Based on its sequence and subunit domain structure, adiponectin is most similar to C1q, a member of the complement-related family of proteins. The molecule shares sequence homology with the type VIII and type X collagens, precerebellin, and the hibernation-regulated proteins hib 20, 25, and 27 (174). The three-dimensional structure of its C-terminal globular domain is remarkably similar to that of TNF-α, even though there is no sequence homology at the primary amino acid level. Based on this observation, Shapiro et al. suggest an evolutionary connection between C1q- like proteins and TNFs, which share some common functions (175).

Adiponectin protein in its most basic form is a homotrimer of three 30 kDa subunits. Trimers associate through disulphide bonds within the collagenous domains of each monomer to form bouquet-like higher order structures that can be found circulating in plasma (176). These higher order structures include a low–molecular weight trimer, a middle–molecular weight hexamer, and high–molecular weight 12- to 18-mer adiponectin of 400-600 kDa (177). Without the collagenous domain, the globular domain of adiponectin still trimerizes, but does not associate into higher-order structures (174). Almost all adiponectin appears to exist as full-length adiponectin in plasma, with a small amount of proteolytically generated globular adiponectin present in plasma (178). The function of this protein however, remains to be determined. Wang et al. suggest that hydroxylation and glycosylation of the lysine residues within the collagenous domain of adiponectin are critical in regulating the formation of its high-molecular weight oligomeric complex and mediating biological functions (179). Serum adiponectin levels are surprisingly high in humans (5-10 μg/mL or 0.01% of serum protein) and mice (0.05%). Males have significantly lower plasma adiponectin levels than females, and this sexual dimorphism develops during pubertal development in relation to serum androgens (180).

Adiponectin exerts its effects by activating the adiponectin receptor (AdipoR1 and AdipoR2). Both receptor isoforms are predicted to contain seven transmembrane domains, but appear to be

33 structurally and functionally distinct from G-protein-coupled receptors (181). In mice, AdipoR1 is largely expressed in skeletal muscle, whereas AdipoR2 is predominantly expressed in the liver. Both receptor subtypes have also recently been detected in the adipose tissue of mice, suggesting adiponectin also acts in an autocrine/paracrine function (182). Globular and full- length adiponectin bind to AdipoR1 and AdipoR2 and mediate activation of AMP-activated protein kinase (AMPK), an enzyme that plays a key role in regulation of carbohydrate and fat metabolism. AMPK phosphorylates numerous target proteins, which results in increases or decreases in the rate of various metabolic pathways (183). By activating AMPK, as well as inducing PPARα expression, adiponectin increases fatty acid oxidation and glucose uptake. AdipoR1 interacts with insulin receptors, thereby enhancing insulin signal transduction (184). More recently, expression of both receptors has been demonstrated in pancreatic β cells in mice and humans, and their expression is increased by exposure to oleate. This suggests that the regulation of adiponectin action, at least in part, can occur by controlling the expression of adiponectin receptors (185).

Adiponectin plays an important role in the modulation of glucose and lipid metabolism in insulin-sensitive tissues. Early studies revealed that adiponectin gene expression was negatively correlated with BMI, and is significantly reduced in the adipose tissues from obese mice and humans (171). It has since been reported that adiponectin levels are also decreased in conditions of insulin resistance and diabetes, and in cardiovascular disease with increasing severity (186). This reduction appears to precede the disorders, suggesting that decreases in adiponectin may play a role in, and be a predictor of the aforementioned pathological states (187). Indeed, Spranger et al. reported that plasma adiponectin concentrations are lower among human subjects who later developed type 2 diabetes than among controls (188). Furthermore, in a study of severely obese Pima Indians, those with high concentrations of adiponectin were 40% less likely to develop type 2 diabetes than those with low concentrations (189).

In vivo and in vitro studies underscore the importance of adiponectin in regulating many metabolic functions. Adiponectin knockout mice show severe insulin resistance and impaired glucose metabolism which can be reversed by adiponectin treatment (190). Conversely, transgenic mice overexpressing adiponectin display improved insulin sensitivity and hypertrophy of adipose tissue (191). Berg et al. observed that adiponectin treatment decreased hepatic glucose production by inhibiting enzymes of gluconeogenesis in obese rats, and also provoked a reduction in blood glucose levels in normal as well as diabetic animals (192). Furthermore, peripheral administration of adiponectin reduces visceral adiposity, increases free fatty acid oxidation, enhances insulin sensitivity and augments glucose uptake by myocytes (193). Recent evidence suggests that adiponectin also acts on the brain to regulate energy balance, since i.c.v. administration of adiponectin decreases body weight by stimulating energy expenditure (194).

Accumulating evidence indicates that adiponectin also possesses anti-inflammatory and anti- atherogenic properties. High blood levels of adiponectin are associated with reduced risk of heart attack, while low levels are found in people who are obese and at increased risk of heart attack. Moreover, plasma adiponectin levels in diabetic subjects with coronary artery disease are lower than in diabetic patients without it (195). Adiponectin exhibits pleiotropic effects on vascular cells, modifying endothelial cell function, proliferation of smooth muscle cells and lipid accumulation of macrophages (196). Mice lacking adiponectin exhibit severe diet-induced

34 insulin resistance and neointimal thickening in response to vascular injury, while overexpression of adiponectin in apoE knockout mice reduces fatty streak formation in aorta (197). In human aortic endothelial cells, adiponectin decreases the surface expression of vascular adhesion molecules known to modulate endothelial inflammatory responses (198). Furthermore, adiponectin has been shown to suppress proliferation and activation of immune cells and the secretion of inflammatory cytokines such as TNFα in the atherogenic process (199). For more information on the role of adiponectin in cardiovascular function, the reader is referred to reviews by Hug and Berg (200).

Circulating adiponectin levels are tightly controlled and remain relatively constant. Numerous hormones and factors are involved in the regulation of adiponectin secretion. Insulin decreases adiponectin levels, whereas thiazolidindiones (TZD) increase the expression of adiponectin, possibly through interaction with the PPARγ response element within the adiponectin promoter Fasshauer, 2003 6285 /id;Iwaki, 2003 6286 /id}. Several factors have been reported to inhibit adiponectin expression and secretion, including catecholamines, glucocorticoids, IL-6, TNFα, GH, and androgens (182;201).

Fasting-induced adipocyte factor (FIAF)

FIAF is a novel adipokine that has been described as a regulator of lipid metabolism and angiogenesis. Discovered in parallel by several groups in 2000, it is also known as PGAR (PPARγ angiopoietin-related protein), Angptl4 (for angiopoietin-like protein 4), or HFARP (for hepatic fibrinogen/angiopoietin-related protein) (202-204).

The FIAF protein is composed of 410 amino acids. It has a predicted molecular mass of approximately 50 kDa, but is glycosylated up to 74 kDa. It bears structural similarities to the angiopoietins, and has thus been classified as an angiopoietin-like protein. The angiopoietins and Angptls encompass a group of about nine proteins that share a common modular structure consisting of an N-terminal signal sequence, a unique region of variable length, a coiled-coil domain, and a large fibrinogen/angiopoietin-like domain. The latter two well-conserved domains comprise the majority of the protein and are involved in oligomerization, giving rise to several high molecular mass species. Oligomeric forms of FIAF involve the formation of intermolecular disulfide bonds. Mutational analysis revealed that cysteine residues at positions 76 and 80 of Angptl4, conserved among mouse, rat, and human, are required to form higher order structures (205). Although angiopoietins and Angptls are indistinguishable structurally, the property that separates the two sets of proteins is the ability of angiopoietins to bind to the Tie2 receptor. Mandard et al. report the presence of two short FIAF isoforms, likely resulting from cleavage of the native 50 kDa protein. These isoforms, deemed S1 and S2, are ~35 kDa and are differentially expressed in several tissues, including adipose tissue and liver. Based on the fact that native FIAF is glycosylated, it has been proposed that FIAF-S1 and FIAF-S2 may represent different glycosylated forms (202;203).

FIAF is predominantly expressed in adipose tissue and liver, but is also found in testes, heart, spleen, lung, kidney, and pituitary (202). In fact, although FIAF is largely recognized as an adipokine, the majority of FIAF appearing in serum may be produced in the liver. Notably, FIAF mRNA expression in adipose tissue and liver, as well as plasma levels of FIAF are increased by

35 fasting and decreased by feeding (202). Notably, with respect to FIAF protein in plasma, native FIAF was found to be elevated after fasting, whereas levels of S1 or S2 did not seem to be affected (206).

The transcriptional regulation of FIAF is complex. Three transcriptional start sites have been identified in murine Angptl4 gene (207). A cis-regulatory element controlling Angptl4 expression was found within intron 3, which binds to both PPARα and PPARγ (206). Both PPARα and PPARγ, as well as hypoxia inducible factor 1α (HIF-1α), activate FIAF transcription (202;204;208). The effect of fasting partially occurs via PPARs. Mice heterozygous for PPARγ or with PPARα deficiency exhibit reduced to undetectable FIAF mRNA levels in liver. Conversely, administration of a potent agonist of PPARα, such as WY-14643 or fenofibrate increases F transcription (202). FIAF appears to be a general target of PPARs, responding to all three PPARs in numerous tissues. In vitro, treatment of NIH 3T3 cells stably expressing PPARγ with its agonist, pioglitazone, causes a significant increase of FIAF mRNA levels in the absence of protein synthesis (209). Recent studies by Yamada et al. suggest that insulin may play a role in the nutritional-mediated regulation of FIAF, as it down-regulated FIAF expression via PI3K/Foxo1 pathway in 3T3-L1 adipocytes (210).

A function for FIAF was discovered in 2002, when Yoshida et al. observed that administration of recombinant FIAF potently raised plasma TGs (211). These authors demonstrated that FIAF treatment inhibited LPL activity in vitro, providing a plausible explanation for its hypertriglyceridemic effect in vivo. Further support comes from studies by Ge and Koster, who reported that either adenovirus-mediated or transgenic overexpression of FIAF also results in severely elevated plasma TG levels under both fed and fasted conditions (212;213). In contrast to the transgenic mice, FIAF -/- mice displayed hypotriglyceridemia and increased LPL activity, with greater effects in the fasted compared with the fed state (213). Mandard and Kersten have concluded that impaired clearance of TG-rich apoB-containing lipoproteins, represented by chylomicrons and VLDL, rather than increased production, is responsible for the observed hypertriglyceridaemia (214). In fact, recombinant mouse FIAF inhibits recombinant human LPL with an IC50 of 14 nM, suggesting direct interactions of FIAF with LPL (213). Mandard et al. recently reported that FIAF overexpression caused a 50% reduction in adipose tissue weight, partly by stimulating fatty acid oxidation and uncoupling in fat (215). In these experiments, FIAF overexpression also increased plasma free fatty acids, glycerol, total cholesterol, and high density lipoprotein (HDL)-cholesterol (215).

Interestingly, the ability of FIAF to elevate plasma TG appears to be dependent on oligomerization, which may stabilize the coiled-coil domain (212). Ge et al. also report that oligomerized FIAF is proteolytically cleaved by an unidentified serum component to release its C-terminal fibrinogen-like domain, which circulates as a monomer. The N-terminal coiled-coil domain which mediates its oligomerization, is still sufficient to form higher order oligomeric structure.

It is not entirely clear which FIAF isoform is responsible for its physiological actions. Only one study has addressed this issue, reporting that full-length is required for activity (212). Notably, in that study, FIAF was cleaved into two fragments in vivo, and levels of the N-term isoform correlated with plasma TG. This suggests that the N-term fragment mediates biological activity.

36 Moreover, this raises the notion that proteolytic processing of FIAF to is a mechanism by which its activity is regulated.

A few recent studies report that FIAF also regulates glucose metabolism. Adenoviral-mediated overexpression of FIAF is associated with a dramatic decrease in basal glucose levels and improved glucose tolerance, while deletion of the FIAF gene or hepatic overexpression did not affect fed or fasted plasma glucose levels (213). Another study found that glucose tolerance was impaired in FIAF transgenic mice fed a high-fat diet (215). Clearly, the role of FIAF in glucose metabolism remains undefined, and more research is necessary.

Expression of FIAF is dramatically up-regulated during hypoxia in both endothelial cells and cardiomyocytes, indicating that FIAF may be involved in angiogenesis (208;216). In support of this notion, recent studies have shown that FIAF has anti-apoptotic activity in vascular endothelial cells and induces a pro-angiogenic response in the chicken chorioallantoic membrane assay (203;216;217). Furthermore, Hermann et al. reported that FIAF protein levels are elevated in stromal fibroblast-like cells within arthritic joints, and that recombinant mouse FIAF promoted endothelial cell survival and formation of tubule-like structures (218).

B. Pancreas

1. Structure

The pancreas is an organ containing two distinct populations of cells, the exocrine cells that secrete enzymes into the digestive track, and the endocrine cells that secrete hormones affecting carbohydrate metabolism into the bloodstream. The exocrine pancreas is a lobular, branched, acinar gland. The secretory cells are grouped into acini, have a pronounced Golgi complex and numerous secretory granules which contain the digestive enzymes. Most are secreted as inactive precursors and become activated once they enter the duodenum.

The pancreas arises from the endoderm as a dorsal and ventral bud which fuse together to form a single organ. Mammals, birds, reptiles, and amphibians have a pancreas with similar histology and mode of development, while in some fish the islets are segregated as Brokkman bodies. Invertebrates do not have a pancreas, but comparable endocrine cells can be found in the gut or brain. In the developing pancreatic buds, the endocrine cells begin to differentiate before the exocrine cells, and coexpression of different hormones by the same cell is often observed at early stages. Development of the pancreas in embryonic life requires a trophic stimulus from the associated mesenchyme (219). In postnatal life, all cell types in the pancreas continue to grow, albeit with a limited capacity.

The endocrine pancreas is comprised of aggregations of cells, dispersed throughout the exocrine pancreas. The cell aggregates resemble ‘islands’ of cells and have been named "islets of Langerhans" after their discovery by Paul Langerhans. Islets of Langerhans are composed of four cell types. The β cells produce insulin, and also an insulin antagonist called amylin, and make up the majority of cells in the islets. The α cells secrete glucagon, the δ cells secrete somatostatin and the PP cells produce pancreatic polypeptide. Some adult cells make peptide YY (PPY) in addition to their primary product (220).

37

In rodents, islets are typically organized as a core of β cells with other cell types at the periphery, while in humans this segregation, although present, is less evident. These islets share specific connections with each other and create a complex and heterogeneous micro-organ. This organization depends on cell adhesion molecules (CAM), which exist on the surface of the endocrine cells (221). The CAMs of the β cells require Ca2+ to be functional, but those of non-β cells are Ca2+-independent. It is estimated that adult humans have approximately 2 million islets, which make up about 2% of the pancreas by weight. In rodents, each islet is composed of 2000– 4000 cells of which 70–80% are β cells, 5% are δ cells and 15–20% are either α cells or PP cells. Endocrine cells producing glucagon, somatostatin, amylin and PPY are present not only in the pancreas, but also elsewhere in the gut (222).

2. β cells and insulin synthesis

This section will focus primarily on the endocrine portion of the pancreas, with an emphasis on β cells, insulin, and the regulation of glucose homeostasis. The islet cells monitor the concentration of glucose in the blood and secrete hormones that exert opposite effects. When blood glucose concentrations rise, the β cells secrete insulin, which reduces blood glucose. Insulin stimulates glucose uptake by a variety of cells and stimulates the conversion of glucose to glycogen in the liver. If glucose levels fall too far, α cells secrete glucagon, which stimulates the breakdown of glycogen to glucose in the liver and therefore increases blood glucose between meals. Optimal control of blood glucose levels depends on delicate changes in insulin production and secretion by the β cells, and on their capacity for a large increase of secretion after meals, which requires large stores of insulin. The insulin-producing potential and functional significance of the β-cells was recognized in 1922 when Frederick Banting and Charles Best were studying pancreatectomized dogs (223).

Insulin mRNA is translated as a single chain precursor called preproinsulin (224). In most species, preproinsulin exists as a single gene, whereas in mice and rats two non-allelic insulin genes are present, called insulin I and insulin II. In adult islets, the non-allelic genes are coordinately expressed and regulated (225). The rodent insulin II and the human genes contain three exons and two introns, while insulin I lacks the second intron. The organization and structure of the insulin has been reviewed in detail (226).

Regulation of insulin transcription relies on the interaction of several sequence motifs in the promoter with a number of ubiquitous and islet specific transcription factors. Insulin genes share a number of conserved DNA motifs in their 5’ flanking region. Notably, E, A, and C1RIPE3b elements seem to be the major determinants of β cell-specific expression of the insulin gene (227). Circulating factors influence the binding of transcription factors to these elements thereby regulating preproinsulin production. Glucose increases binding of Pdx-1 to an A element in the promoter, thereby upregulating transcription (228). Certain fatty acids, such as palmitate, inhibit preproinsulin transcription. This effect appears to be mediated by direct inhibition of insulin promoter activity, possibly via the production of ceramide (229). Prolonged exposure to fatty acids can result in lipotoxicity and β cell death. Interestingly, a recent study suggests that suppressor of cytokine signaling-3 (SOCS3) represents a transcriptional inhibitor of

38 preproinsulin gene expression, which is induced by leptin through Jak/Stat3/5b signaling in pancreatic beta-cells (230).

Proinsulin consists of three domains: an amino-terminal B chain, a carboxy-terminal A chain and a connecting peptide in the middle known as the C peptide. Within the endoplasmic reticulum, proinsulin is exposed to several specific endopeptidases which remove the C peptide. The A and B chains are bound together by disulfide bonds in the mature form of insulin. Free C peptide and insulin are packaged in the Golgi into secretory granules which accumulate in the cytoplasm. When the β cell is appropriately stimulated, both insulin and the C peptide are secreted from the cell by exocytosis and diffuse into islet capillary blood.

3. Insulin secretion and the role of glucose

Pancreatic β cells function as glucose sensors with the crucial task of perfectly adjusting insulin release to blood glucose levels. In the consensus model of glucose-stimulated insulin secretion, glucose equilibrates across the plasma membrane via facilitated diffusion through GLUT2, and is phosphorylated by glucokinase (GK) to produce glucose-6-phosphate. The thermodynamic, kinetic and molecular genetic characteristics of GK are ideally suited for its glucose sensor function: the reaction is virtually irreversible; there is no direct feedback inhibition by any of the reaction products; GK shows cooperative kinetics with glucose; and GK has an allosteric activator site (231). As the pacemaker of glucose metabolism in β-cells, GK determines the generation of factors that couple metabolism to the increase of insulin secretion.

Increased levels of glucose-6-phosphate rapidly initiate glycolysis (232). Subsequently, mitochondrial metabolism generates ATP via oxidative phosphorylation, leading to a rise in the ATP/ADP ratio. This leads to a graded inhibition of the K+ efflux, primarily due to the lowering of ADP, a potent activator of the channel (233). The reduction of outward K+ current 2+ 2+ subsequently triggers Ca influx through L-type voltage gated Ca channels (234). The resulting rise in cytoplasmic free Ca2+ leads to activation of exocytotic machinery.

+ Another pathway of glucose-stimulated insulin secretion now being recognized is the K ATP channel-independent but Ca2+-dependent pathway (235). The underlying messengers involved in + this pathway are not yet known. Several hypotheses regarding second messengers in K ATP channel-independent insulin secretion are currently being investigated, including 1) glucose mediated production of glutamate, which may sensitize the secretory machinery to Ca2+, 2) glucose induced activation of PKC, 3) activation of HSL, which may increase long-chain acyl- + CoA and diacylglycerol, and 4) changes in ATP concentrations that may facilitate the K ATP channel-independent pathway (e.g. by activation of PKA, which may constitute an ATP sensor in exocytosis) (236).

In addition to glucose, numerous factors affect insulin secretion. For instance, acetylcholine and cholecystokinin stimulate secretion via the phospholipase C-PKC pathway (237;238). Gastrointestinal inhibitory peptide also stimulates insulin secretion by altering cAMP levels (239). Leptin primarily acts via central mechanisms to suppress insulin secretion, but appears to exert direct effects on pancreatic cells under certain conditions. For instance, while leptin alone had no effect on insulin release from INS-1 cells under normal culture conditions, it inhibited

39 secretion when cAMP levels were elevated (240). There are some inconsistencies in the literature, as one study reported that leptin suppressed insulin secretion from isolated rat islets in basal glucose media (5.5 mM), but not under glucose stimulated conditions (11.1 mM), while another study reported that leptin was a potent inhibitor of basal and glucose-induced insulin secretion (241;242). Interestingly, in islets isolated from ob/ob mice, leptin had no effect on insulin secretion in the presence of low or high levels of glucose (2.8-20 mM) (243). The direct effects of adiponectin on insulin secretion also appear to vary under different glucose concentrations. In islets isolated from normal mice, physiological doses of adiponectin had no significant effect on insulin secretion. In contrast, in those from mice rendered insulin resistant by high-fat diet, adiponectin inhibited insulin secretion at low glucose levels, but augmented insulin secretion at high glucose levels (244). Taken together, it is obvious that numerous factors contribute to the regulation of insulin production, notably by hormones whose own secretion is dependent on energy status.

When blood glucose levels decrease to normal physiological values, insulin release slows or stops. A drop to low levels induces the release of hyperglycemic hormones, predominantly glucagon, which increases the release of glucose into the blood from cellular stores, primarily glycogen stored in liver cells, which will be described in the next section.

The proper regulation of β cell mass is crucial. Glucose, as well as several hormones, affect β cell replication, size, apoptosis, and under certain conditions, neogenesis from progenitor cells. Failure to adapt to changes in body mass, pregnancy, reduced insulin sensitivity of peripheral tissues, or tissue injury may lead to the development of chronically elevated blood glucose, or diabetes (245). In humans, a critical reduction in beta-cell mass is primarily responsible for the development of type 1 diabetes, which is characterized by lack of insulin production. Transplantation studies have demonstrated that the diabetic metabolic defects can be ameliorated in type 1 patients by restoring a functional beta-cell mass (246). Type 1 diabetes usually develops due to an autoimmune disorder, but the specific cause of this remains unknown. Genetic factors certainly play a role, with some people being more prone to develop diabetes than others. Type 2 diabetes, characterized by low or absent insulin sensitivity, varies widely among populations and usually develops during mid- to late life. Common symptoms include weight gain and often obesity. Insulin levels in these patients are usually normal or higher than average but peripheral cell response is blunted or absent. This lack of insulin action results in higher than normal blood glucose levels.

Pregnancy induces profound alterations in maternal metabolism in the presence of increasing fetal demand for more fuel. This demand is met through increased caloric intake, an elevated insulin secretory response, insulin resistance in some maternal tissues, and increased lipid metabolism. The pancreas plays a major role in these adaptations. During pregnancy, β cells undergo several structural and functional changes in response to the increased demand for insulin. These include 1) increased glucose-stimulated insulin secretion due to a lowered threshold for glucose, 2) increased insulin synthesis, 3) increased β cell proliferation and hypertrophy, 4) increased gap-junctional coupling among β cells, 5) increased glucose metabolism, and 6) increased cAMP metabolism (247).

40 The increase in β cell division and enhanced glucose sensitivity of insulin secretion are the most notable changes that occur during pregnancy (248). The lowering of the threshold for glucose stimulated insulin secretion is the primary mechanism by which β cells release significantly more insulin under normal blood glucose concentrations. Increased β cell division leads to an increase in islet mass that contributes to the ability of the islets to respond to the increased need for insulin. Indeed, at the end of pregnancy in rats, the beta-cell mass is ~2.5-fold higher than in nonpregnant females (249).

Although the hormonal changes which occur during pregnancy are complex, it appears that lactogenic hormones are sufficient to induce all of the up-regulatory changes that occur in islets during pregnancy. This will be discussed in the final chapter of this literature review.

C. Liver

The liver plays a major role in blood glucose homeostasis by maintaining a balance between glucose uptake, storage, and production. It is also an important site of lipid metabolism, where TG, phospholipids, cholesterol, and apolipoproteins are packaged into VLDLs. In addition, the liver is the main organ responsible for catabolism and detoxification.

Hepatocytes make up 60-80% of the cytoplasmic mass of the liver. These cells are involved in protein synthesis, storage and transformation of carbohydrates, synthesis of cholesterol, bile salts and phospholipids, and detoxification. Hepatocytes are organized into plates separated by vascular channels called sinusoids, an arrangement supported by a collagen type III network. The sinusoids display a discontinuous, fenestrated endothelial cell lining. The endothelial cells have no basement membrane and are separated from the hepatocytes by the “Space of Disse,” which enables intercellular exchange of metabolites. Kupffer cells, specialized macrophages that form part of the reticuloendothelial system, are scattered among endothelial cells. Hepatic stellate cells, also known as Ito cells, are pericytes found in the perisinusoidal space. These cells store vitamin A in lipid droplets and produce extracellular matrix and collagen.

The liver plays a role in several aspects of carbohydrate metabolism, including gluconeogenesis (the formation of glucose from amino acids, lactate or glycerol), glycogenolysis (the formation of glucose from glycogen), and glycogenesis/glyconeogenesis (the formation of glycogen from glucose).

1. Glucose metabolism

The major site of daily glucose consumption is the brain via aerobic pathways. Most of the remainder is utilized by skeletal muscle, heart muscle, and erythrocytes. The body obtains glucose either directly from the diet, or from amino acids and lactate via gluconeogenesis. Glucose obtained from these two primary sources either remains soluble in the body fluids or is stored in a polymeric form, glycogen. Glycogen is considered the principal storage form of glucose and is found mainly in liver and muscle, with kidney and intestines adding a minor storage capacity. With up to 10% of its weight as glycogen, the liver has the highest content of any body tissue. Muscle has a much lower amount of glycogen per unit mass of tissue, but since the total mass of muscle is much greater than that of liver, total glycogen stored in muscle is

41 about twice that of liver. Stores of glycogen in the liver are considered the main buffer of blood glucose levels.

Glycogen is a polymer of glucose residues linked mainly by α-1,4 glycosidic linkages, with α1,6 linkages occurring at branch points. Synthesis of glycogen from glucose is carried out by glycogen synthase (250). This enzyme utilizes UDP-glucose as one substrate and the non- reducing end of glycogen as another. The activation of glucose is carried out by UDP-glucose pyrophosphorylase, which exchanges the phosphate on C-1 of glucose-1-phosphate for UDP. The energy of the phospho-glycosyl bond of UDP-glucose is utilized by glycogen synthase to catalyze the incorporation of glucose into glycogen. Because glycogen synthase can only add to an existing chain, glycogenin acts as the “primer" to begin this mechanism (251). Glycogenin can catalyze the addition of glucose to itself by binding the first UDP-glucose to its active site. It then adds eight additional glucose residues, at which point glycogen synthase takes over.

Glycogen synthase activity is negatively regulated by phosphorylation. The kinases responsible for this covalent modification are controlled by the second messengers generated by different hormones (252). For instance, activators of the “fight or flight” mechanism, such as epinephrine, initiate signaling pathways that inhibit glycogen synthase. Activation of glycogen synthase requires dephosphorylation, which is carried out predominately by protein phosphatase-1 (PP-1). When glucose levels rise, insulin attenuates the activity of kinases such as PKA or glucose synthase kinase-3 and activates PP-1, thereby increasing the activity of glycogen synthase (253).

When blood glucose levels drop, glycogen is degraded into glucose subunits in a process termed glycogenolysis. This occurs through the action of glycogen phosphorylase, which uses inorganic phosphate (Pi) and removes single glucose residues from α-(1,4)-linkages within the glycogen molecules. The product of this reaction is glucose-1-phosphate. The advantage of this reaction is that glucose is produced in an activated state without the use of ATP, and the concentration of Pi in the cell drives the equilibrium of the reaction the favorable direction since the free energy change of the standard state reaction is positive. Glucose-1-phosphate is converted to glucose-6- phosphate by phosphoglucomutase. Since the cell membrane is impermeable to glucose-6- phosphate, this process effectively traps glucose inside the cell. In liver, kidney, and intestine, glucose-6-phosphate is converted to glucose by glucose-6-phosphatase, allowing glycogenolysis to generate free glucose for distribution to other tissues. This does not occur in skeletal muscle since the cells of this tissue lack glucose-6-phosphatase. Therefore, any glucose released from glycogen stores of muscle will be oxidized in the glycolytic pathway.

Phosphorylation of glycogen phosphorylase is mediated by phosphorylase kinase, which is activated by PKA. When blood glucose levels are low, α-cells secrete glucagon, which binds to G protein-coupled receptors on hepatocytes. The activated PKA phosphorylates glycogen phosphorylase and enhances glycogenolysis. Calcium ions indirectly lead to activation of glycogen phosphorylase, allowing neuromuscular stimulation by acetylcholine to increase glycogenolysis in the absence of receptor stimulation.

Buffering blood glucose by the liver is made possible by GK, which phosphorylates glucose to glucose-6-phosphate, and its activity is altered by blood glucose over the physiological range. Thus, at low blood glucose levels, the liver does not compete with other tissues for the available

42 glucose supply. Conversely, at high blood glucose levels, glucokinase converts glucose to glucose-6-phosphate, trapping it in hepatocytes and promoting the formation of glycogen.

De novo synthesis of glucose from precursors such as lactate, amino acids, and glycerol is an important mechanism for providing the organism with glucose in times of starvation. The production of glucose from other metabolites is necessary for use as a fuel source by the brain, testes, and erythrocytes since glucose is the sole energy source for these organs. This process takes place primarily in the liver, and to a lesser extent in the kidney (254). Synthesis of glucose from three and four carbon precursors is essentially a reversal of glycolysis, and utilizes many of the same enzymes. Three reactions of glycolysis have such a large negative ΔG in the forward direction that they are essentially irreversible, and are bypassed in gluconeogenesis by specific enzymes.

Lactate is a predominate source of carbon atoms for gluconeogenesis. Formed during anaerobic glycolysis in skeletal muscle, lactate is released into the bloodstream and transported to the liver where it is converted first to pyruvate and then to glucose. This process is termed the Cori cycle (255). The glycerol backbone of lipids can also be used for gluconeogenesis. This requires phosphorylation to glycerol-3-phosphate by glycerol kinase and dehydrogenation to dihydroxyacetone phosphate by glyceraldehyde-3-phosphate dehydrogenase (256). The glycerol backbone of adipose tissue stored TG is used as a gluconeogenic substrate since adipocytes express low levels of glycerol kinase. All amino acids except leucine and lysine can be degraded to TCA cycle intermediates, allowing their carbon skeletons to be converted to pyruvate, oxaloacetate, or their precursors (257). Pyruvate can be utilized by the gluconeogenic pathway. When glycogen stores are depleted in muscle during exercise and in liver during fasting, catabolism of muscle proteins to amino acids constitutes the major source of carbon for maintenance of blood glucose levels. Tight regulation of gluconeogenesis is necessary to ensure proper buffering of glucose levels.

While de novo synthesis of glucose provides the organism with energy in times of starvation, gluconeogenesis is dispensable and therefore shut off when glucose is directly available from external resources. When glucose is delivered to the liver in sufficient quantities that glycogen stores are re-established, excess glucose can be converted in the liver into triglycerides through de novo lipogenesis.

2. Lipid metabolism

In addition to its role in maintaining glucose homeostasis, the liver is an important site of lipid metabolism, where TG, phospholipids, cholesterol, and apolipoproteins are packaged into very- low density lipoproteins (VLDL).

Long chain fatty acids can be synthesized from acetyl-CoA produced by the catabolism of excess glucose. The metabolic fates of LCFA in the liver are 1), the esterification into TG, and to a lesser extent, into phospholipids and cholesteryl esters 2) complete oxidation to CO2 generating ATP molecules or incomplete oxidation to acetate and ketone bodies (258).

43 TG produced in the liver must be transported to other tissues for use. In contrast to water-soluble fuels such as glucose or ketone bodies, the use of lipids, which are largely insoluble, as an energy source for tissues has required the development of complex structures for their transport through the aqueous plasma. Hence, they are transported by the circulation as components of lipoproteins; globular micelle-like particles that consist of a nonpolar core of TG and cholesteryl esters surrounded by an amphiphilic coating of protein, phospholipids, and cholesterol. Lipoproteins are classified into five broad categories on the basis of their function and physical properties: 1) chylomicrons, which transport exogenous TG and cholesterol from the intestines to other tissues, 2) high density lipoproteins (HDL), which transport endogenous cholesterol from tissues to the liver, and 3-5) very low density lipoproteins (VLDL), intermediate density lipoproteins (IDL), and low density lipoproteins (LDL), a group of related particles that transport endogenously synthesized TG and cholesterol.

The liver produces and secretes VLDL into the bloodstream where they function as the body's internal transport mechanism for lipids. This allows for efficient mobilization of a powerful source of energy either for storage in adipose tissue or for utilization, predominantly by muscle tissues. VLDL are complex particles consisting of a core of neutral lipids (mostly TG) surrounded by a monolayer of amphipathic lipids such as phospholipids and unesterified cholesterol. Apolipoprotein B-100 (apoB-100), a protein which forms the structural framework for the VLDL, is bound to the surface (259). The assembly of VLDL occurs in two major steps (260). The first step is the co-and post-translational lipidation of apoB, forming pre-VLDL in the rough endoplasmic reticulum (261). The partially lipidated apoB (the pre-VLDL) is transported out of the endoplasmic reticulum before it can acquire the major amount of lipids to form VLDL. In the second step pre-VLDL is converted to bona fide VLDL in a smooth membrane compartment. This step depends on ADP-ribosylation factor 1 and its activation of phopholipase D. The addition of the major amount of lipids may involve the production of an apoB-free particle containing the core lipids, which could fuse with the pre-VLDL in the second-step compartment. When VLDL is completed it is allowed to transfer into the later part of the Golgi apparatus to eventually be secreted in an incomplete nascent state.

Nascent VLDL circulates in the blood and acquires apolipoprotein C-II (apoC-II) and apolipoprotein E, which are donated from HDL, resulting in the formation of a mature VLDL (262). As described earlier, the TG component of VLDL (and chylomicrons), are hydrolyzed through the action of LPL, which is activated by apoC-II. VLDL are degraded into VLDL remnants, which appear in the circulation as IDL, and then as LDL. In the transformation from VLDL to LDL, all proteins but apoB are removed and much of the cholesterol is esterified by HDL. LDL subsequently transfers cholesterol either back to the liver or to other tissues. This is achieved through the endocytosis of LDL particles in a process mediated by the LDL receptor, a cell-surface transmembrane glycoprotein, which specifically binds apoB-100 and apoE.

In summary, the liver is an essential metabolic organ, functioning to maintain proper levels of nutrients in the blood for use by the brain muscles, and other tissues. The liver is uniquely positioned to carry out this task because the nutrients absorbed by the intestines are released into the portal vein, which drains directly into the liver.

D. Brain

44

The brain plays a vital role in metabolism, integrating peripheral metabolic, endocrine, and neuronal signals that reflect current energy status to coordinate a modulating effect on behavioral patterns and peripheral metabolism according to acute and chronic requirements.

There is a tendency to simplify the regional contribution of brain areas to metabolic regulation. This viewpoint has its advantages, as it allows easier conceptualization of the involvement of brain regions, as well as neuropeptides and their receptors, in energy metabolism. However, focusing on specific areas of the brain, such as the arcuate nucleus, is a dangerous tactic, as it is easy to get too involved in the details of the system rather than seeing it as a whole, and nearly impossible to integrate all contributing factors and pathways. Thus, when brain structures and their recognized functions are discussed in this section, the reader is cautioned not to consider this a set blueprint for the only brain regions/mechanisms involved in metabolic regulation. Instead, this section should be viewed as a broad discussion, which introduces a few major pathways that pertain to the satiety/energy expenditure signals studied in this thesis.

The hypothalamus is a specialized area in the CNS that integrates the control of energy homeostasis. Discrete nuclei within the hypothalamus respond to changes in energy status by altering the expression of specific neuropeptides, which cause changes in food intake and energy expenditure. Nuclei directly associated with metabolic regulation are the PVN, arcuate nucleus, lateral hyptothalamus-perifornical region, and ventromedial (VMH) and dorsomedial (DMH) hypothalamic nuclei (263). The arcuate nucleus sits in the mediobasal hypothalamus, between the third ventrical and the median eminance, where the portal vascular system functions to transport neuroendocrine releasing factors to the anterior pituitary. Here it is exposed to factors both in the blood and the cerebrospinal fluid. Though not classically viewed as such, the arcuate nucleus may sometimes function as a circumventricular organ, existing functionally outside the blood-brain barrier (264).

Among the hypothalamic neuropeptide systems regulating feeding, melanocortins play a prominent role (265). The central melanocortin system modulates energy homeostasis through the anorectic actions of α-melanocyte stimulating hormone (α-MSH), and the endogenous orexigenic effects of agouti-related protein (AgRP) and neuropeptide Y (NPY).

α-MSH belongs to a group of hormones that includes β-MSH and γ-MSH. These peptides are all cleaved products of the precursor molecule proopiomelanocortin (POMC), which is synthesized by corticotrope cells of the AP, melanotrope cells of the intermediate pituitary, the arcuate nucleus, and smaller populations of neurons in the dorsomedial hypothalamus and brainstem. In mice there are approximately 3,000 hypothalamic POMC neurons, most of which also express the anorectic peptide Cocaine- and Amphetamine Related Transcript (CART). POMC- and CART- (POMC/CART) cell bodies are found in the arcuate nucleus and periarcuate area (266). Those neurons located in the arcuate nucleus send a dense bundle of fibers to several regions of the brain, notably within the hypothalamus, the PVN, and the perifornical region (266). These hypothalamic POMC neurons also send two descending sets of projections to the brainstem (267).

45 Melanocortins exert their effects by binding to members of a family of G-protein coupled melanocortin receptors (MC1 to MC5) (268). α-MSH is a potent agonist for MC1, as well as MC3 and MC4, which are expressed within the CNS. The distribution of MC4 is consistent with the proposed roles of central melanocortin systems in feeding and autonomic regulation (269). POMC or MC4 deficiency produces obesity in mice and humans, while mice heterozygous for MC4 are moderately obese (270). Administration of synthetic antagonists for MC4 receptors increases food intake (271). On the other hand, central administration of α-MSH in wild-type mice results in potent reductions in food intake and significant loss of body weight (272). Although MC4R involvement in the regulation of feeding is well established, the role of MC3 remains less. Relatively selective MC3 agonists do not alter food intake, and unlike MC4 expression which is influenced by energy status, MC3 expression is not (273).

MC3 are MC4 are bound not only by α-MSH, but also by AgRP, a neuropeptide that is coexpressed with NPY in neurons within the arcuate nucleus (274). AgRP has little intrinsic signaling activity, and instead functions primarily by inhibiting α-MSH binding (275). AgRP is upregulated by fasting, and the wide distribution of AgRP terminals suggest that it participates in the regulation of food consumption through several hypothalamic structures, including the PVN, arcuate, dorsomedial, and lateral hypothalamic nuclei, as well as the amygdala, an area implicated in emotional aspects of feeding behavior (276). AgRP-immunoreactive fibers occur primarily in a subset of the same hypothalamic and septal brain regions containing dense POMC innervation, with the densest fibers found in the paraventricular hypothalamus, DMH, and posterior hypothalamus and septal regions around the anterior commissure (274).

The best-documented aspect of AgRP action is its profound stimulation of food intake, via inhibition of orexigenic melanocortin signals. Transgenic mice overexpressing AgRP exhibit severe obesity, hyperinsulinemia, late-onset hyperglycemia, and pancreatic islet hyperplasia (277). These effects of AgRP are proposed to be mediated predominantly by the brain, as AgRP stimulates hyperphagia when administered i.c.v. (278). In addition to alterations in feeding behavior, recent data indicate that AgRP also acts to suppress energy expenditure. For example, i.c.v. administration of N-terminal fragments of AgRP increases body weight and fat content in the absence of hyperphagia (279). Conversely, reduction of AgRP by RNA interference increases energy expenditure and loss of body weight without changes in food intake (280). AgRP also influences neuroendocrine function by regulating the HPA axis. In rhesus monkeys i.c.v. injection of AgRP stimulated ACTH, cortisol, and PRL release and enhanced the ACTH response to IL-1β, suggesting a role in inflammation responses (281).

NPY is one of the most abundant and widely distributed neuropeptides within the nervous system and is also a potent stimulator of feeding and suppressor of energy expenditure (282). NPY is coexpressed with AgRP in neurons of the arcuate nucleus, and exerts its effects through NPY receptors. The numerous G-protein receptor Y-receptors mediate their response by inhibiting the accumulation of cAMP, although Y1 and Y2 have also been shown to increase intracellular Ca2+ concentrations.

The interplay between the POMC and AgRP/NPY neurons regulates both food intake and body weight. However, while deletion of the NPY gene partially suppresses obesity in ob/ob mice, normal mice with combined deletion of both NPY and AgRP suffer only limited deficits in

46 feeding or body weight (283;284). These and other observations indicate that the systems regulating food intake and body weight have high redundancy (285). Clearly, central control of appetite and energy balance involves widely distributed neural systems in the brainstem, cerebral cortex, olfactory areas, and elsewhere. Moreover, the collection of circuits regulating food intake and energy homeostasis is unique in having the capability of sensing signals from a staggering array of hormones, nutrients and afferent neural inputs.

Leptin and insulin, both of which reflect the total amount of fat in the body, have a significant influence on the activity of POMC and AgRP/NPY neurons. Receptors for both hormones are located throughout the brain, notably in hypothalamic POMC and AgRP/NPY neurons (286). Changes in the level of either insulin or leptin within the arcuate nucleus cause alterations in food intake and body weight. Increased insulin or leptin activity results in the stimulation of POMC and inhibition of NPY/AgRP, which leads to decreased food intake, increased energy expenditure and weight loss. Conversely, decreased insulin or leptin activity results in the opposite effects. Arcuate NPY neurons also express the receptor for ghrelin, an appetite stimulatory factor produced by the stomach (287).

Circulating lipids also act as signaling molecules informing about metabolic status. I.c.v. administration of oleic acid (OA) inhibits food intake in rats (288). These effects are exerted on the arcuate nucleus, as AgRP and NPY expression was decreased after OA treatment. POMC expression was not affected by OA treatment suggesting that POMC actions do not mediate LCFAs anorectic effects.

Glucose sensing in the brain is also plays a role in energy balance. In 1969, Oomura et al. reported that populations of neurons in the VMH and in the lateral hypothalamus increase their firing rates in response to the application of glucose (289). These glucose-sensitive/responsive neurons are likely to be responsible for the effects of glucose deprivation to induce feeding. In fact, meal onset was historically thought to be a reflexive response due to the reduction of glucose utilization by these sensor cells (290).

Whereas forebrain structures such as the arcuate nucleus are critical sites for sensing adiposity- related input such as leptin and insulin, information provided by meal-related signals is conveyed primarily to hindbrain areas such as the nucleus of the solitary tract, with afferent vagal nerve fibers playing a central role in this process (291). For example, the gastrointestinal hormone cholecystokinin (CCK) is released in response to nutrient stimulation to signal fullness (285). Additionally, entry of food into the stomach and proximal small intestine activates stretch and mechano-receptors, which transmit signals to the hindbrain where integration of this visceral input occurs. Nutrients arriving via the portal vein may also trigger vagal afferent signals from the liver. Notably, these short-term signals by themselves do not produce sustained alterations in energy intake and body adiposity.

While there are certainly other circuits and factors that contribute to the regulation of metabolic homeostasis by the brain, they can not all be included in this review. Instead I have described those brain regions best associated with food intake/energy expenditure, and the pathways most relevant to the factors studied in this thesis. On that note, it can be summarized that signals of long-term energy stores and short-term fluctuations in food intake are integrated in the

47 hypothalamus and brain stem. Important neuropeptide signals such as NPY, AgRP, and the melanocortins are released and influence activity of diverse circuits within other hypothalamic nuclei, which signal using a wide range of transmitter systems. This homeostatic process results in subsequent changes in appetite, behavior, and energy expenditure.

Section II. Integrative regulation of metabolic homeostasis

This section will integrate the organs and systems presented in the previous sections, hopefully giving the reader a better appreciation of the complex interactions of the numerous factors and pathways involved in regulating metabolic homeostasis. It would be very easy to get wrapped up in a lengthy, detailed description of any one energy signal or pathway, but that would dilute the focus and flow of this section. Moreover, it will not be possible to discuss every factor or pathway that contributes to overall metabolic homeostasis. Instead, I will focus on those systems that were discussed in this thesis. Some points from Section I will be repeated, while some previously unmentioned information may be presented to support the issues being addressed.

A. Overall Considerations

Energy homeostasis in higher organisms is under the control of an integrated system that has the capacity to rapidly respond to metabolic changes. This system relies on the functions of numerous tissues as well as their interactions with one another to orchestrate the seamless transition between widely varying states, including the fed state, fasting, and exercise. Proper control of these systems relies upon the detection and integration of signals (derived from adiposity, satiety, and nutritional signals) reflecting long- and short-term energy stores and fluxes, and their interaction with a multitude of additional factors (some not discussed in this thesis) including hormonal, as well as learning, social stress, and others.

Perhaps the best way to describe the complex integrative nature of metabolism is to detail the functions performed by each organ and the signals that interconnect them during various physiological states. First, it should be noted that the maintenance of energy balance involves coordinated changes in energy intake and expenditure, and these two limbs of energy balance are physiologically linked, most importantly through brain circuits. Indeed, the brain is the major site were afferent energy signals converge and are integrated to determine the direction and magnitude of efferent responses. The various afferent inputs that the brain uses to adjust food intake and energy metabolism can be broadly subdivided into two groups: those that communicate information pertaining to body energy stores and to the amount of energy consumed over a more prolonged period of time, and those signals that are generated acutely in response to nutrient ingestion.

Second, although there are many peripheral signals that can contribute to energy balance, short- term and long-term food intake and energy balance are regulated through distinct, but interacting, mechanisms. Short-term regulatory signals have a markedly different function than the long-term regulators of energy homeostasis. Among afferent signals reflecting the size of body adipose mass, insulin and leptin are the best studied and understood, and both appear to be required by the CNS for the control of food intake, body weight, and metabolic homeostasis. Although leptin and insulin are produced at different sites, both signals are slowly transported into the CNS, over

48 a period of hours after circulating concentrations increase where they exert relatively long-lived inhibitory effects on food intake while increasing energy expenditure and weight loss in response to an increase in body fat (292). By comparison, signals responding to recently ingested nutrients are more varied and function primarily on a meal-to-meal basis to control gastric emptying and the timing of meal initiation and termination (293).

Long-term energy signals ensure that energy homeostasis is maintained and that body weight and adiposity remain relatively constant. Indeed, there is considerable evidence that body weight and fat content are well regulated. In adult animals and humans, body weight remains within a relatively narrow range, despite large day-to-day fluctuations in the amount of food consumed. Although major changes of body adiposity can be induced in humans and animals by restricting energy expenditure or by overfeeding, body weight and adiposity return very close to baseline levels when ad libitum feeding is resumed. A notable theory has been put forward by Woods et al., who believe that rather than there being a set point for body weight, “there is a range of body weight that an individual is willing to accept and defend” (286).

When considering the integration of adiposity and satiety signals, it is important to appreciate that the negative feedback circuits related to body fat and meal ingestion can easily be overridden by situational events, such as sight, smell and perceived palatability of some desirable food (286). On the other hand, certain factors such as stress can curtail energy intake even when food is available and needed. Given these exceptions it becomes clear that trying to relate energy intake from an individual meal to recent energy expenditure or to fat stores is futile, at least in the short term. Instead, the influence of homeostatic signals becomes apparent only when intake is considered over longer intervals. In other words, what counts is the cumulative effect. For example, obesity develops during a dynamic phase during which fat balance remains positive for a prolonged period of time. However, this process does not necessarily occur on consecutive days: positive fat balance on one day may be partially compensated (or not) by negative fat balance on subsequent days.

Adiposity signals can also control food intake by changing the sensitivity to satiety signals. Indeed, an important interaction of insulin and leptin with CCK has been demonstrated in several studies (294;295). For example, a dose of peripheral CCK that reduces short-term food intake by more than 50% in fed rats is ineffective at reducing meal size in rats in which circulating leptin levels are reduced by fasting. However, when the fall of leptin in fasted rats is prevented by administering leptin, the ability of CCK to decrease food intake in fasted animals is restored. Similarly, energy intake decreases by more than 50% with the combination of insulin plus CCK, suggesting that sensitivity to the effect of CCK to induce satiety is enhanced by insulin's action in the brain. Insulin increases the sensitivity to other satiety signals, including amylin and neuropeptide corticotropin-releasing hormone (286).

In conclusion, the integration of peripheral signals that regulate energy homeostasis in the brain is complex, and not yet entirely understood. However, as stated by Woods et al., the integration of satiety signals with other signals that influence short-term energy intake occurs in vagal afferent fibers and continues into the hindbrain, where meal size is ultimately determined. Concurrently, the hypothalamus receives signals related to fat stores as well as information on

49 meals from the hindbrain, thereby monitoring ongoing metabolism directly and providing it with the capacity to integrate multiple signals that determine ingestion (286).

B. Energy storage and utilization

Food supplies enough energy to support the many functions of the body and this energy comes from fats, carbohydrates, and proteins. Of the three, fat is the most concentrated source of energy because it furnishes more than twice as much energy as protein or carbohydrate (296).

Efficient storage of ingested nutrients and the appropriate regulation of blood glucose levels after a meal are hallmarks of healthy carbohydrate and lipid metabolism. This involves coordinated regulation of the major peripheral organs of carbohydrate and lipid flux: pancreas, liver, skeletal muscle, and adipose tissue. An early event that initiates coordinated patterns, and begins before the meal-related nutrient load has impacted on the gastrointestinal tract and bloodstream, is nervous system-driven insulin release, which is followed by a more sustained release driven by the rising blood glucose (297). As ingested food interacts with the stomach and intestine, short- term signals from the gut function to coordinate and optimize the digestive process, as well as initiate acute negative energy balance, i.e. cessation of food intake.

When a typical meal is ingested, the influx of glucose could potentially increase the plasma glucose concentration eightfold. This does not happen because coordinated mechanisms come into play to increase the disposal of glucose from plasma and to suppress the entry into the circulation of endogenous glucose. By these means, the rise in plasma glucose concentration and the exposure of tissues to hyperglycemia are minimized. Insulin plays a predominant role in this process, activating a variety of metabolic pathways including stimulation of glucose uptake in most peripheral tissues. In the liver, insulin regulates several pathways to increase net hepatic glucose uptake, including suppression of glucose-producing pathways as well as stimulation of anabolic pathways of glucose disposal. These actions collectively restrict post-prandial increases in plasma glucose concentrations. In addition, when glucose concentrations are elevated, the liver has the ability to synthesize lipids de novo through the lipogenic pathway, which could be considered as a metabolic safety valve for storage of carbohydrate energy present in excess of carbohydrate oxidative needs. Deposition of lipid reserves can then be thought to be part of a longer-term feeding strategy.

Proper buffering of TG levels is also an important component of maintaining metabolic homeostasis. Absorption of a typical meal could in principle raise the plasma TG concentration by ten-fold. However, after ingesting a normal meal, plasma TG concentration rises less than 100% in healthy human subjects (298). Similar to carbohydrate metabolism, there are mechanisms that 'buffer' the influx of TG into the circulation and prevent the exposure of tissues to excessive TG flux. In the fed state, adipose tissue is the major site for uptake of dietary TG, utilizing the pathway mediated by LPL−hydrolysis of circulating lipoprotein-TG. This is followed by uptake of the fatty acids, and esterification to glycerol 3-phosphate. Both activation of adipose tissue LPL and esterification are stimulated by insulin. Inhibition of lipolysis by insulin during the postprandial period and an efficient acylation of newly arrived FA inside adipocytes help to create a gradient for FA uptake in adipose tissue (299). As a result, over 90% of LPL-released FA is trapped by adipocytes after feeding (300). The efficiency of uptake is

50 evidenced by the fact that postprandial plasma FFA levels are actually lower than during fasting. This reduction in plasma FA results both from insulin-mediated inhibition of HSL-mediated lipolysis and the facilitated uptake of chylomicron or VLDL-generated FA.

Dietary composition significantly affects the energy storage systems activated. In the case of high carbohydrate diets, fuel storage is dependent on energy balance (301). Under isoenergetic conditions of feeding, when energy balance is in equilibrium, the conversion of carbohydrates into fat does not provide any net storage of fat to the body. The fat synthesized by de novo lipogenesis in some tissues is balanced by simultaneous fat oxidation in others. Under carbohydrate overfeeding conditions, i.e., a persisting positive energy balance, can lead to positive fat balance via two mechanisms; increased carbohydrate oxidation inhibits fat oxidation (an anti-lipolytic action of insulin) contributing to a positive fat balance. Subsequently, the continuous increase in carbohydrate balance results in an increase in glycogen stores, which become progressively saturated. Substantial rates of carbohydrate conversion into fat are induced only when the glycogen stores are first enlarged and carbohydrate overfeeding is maintained.

Consumption of a high-fat meal produces significantly smaller increases in plasma insulin than high-carbohydrate meals, resulting in reduced circulating leptin concentrations over a 24 hr period (302). Moreover, failure to elevate hypothalamic insulin content following a high-fat meal results in the lack of an additional inhibitory satiety signal. Chronic consumption of a high fat diet can impair brain insulin transport, and the impairment is predictive of weight gain in response to the high fat feeding (303). Together, these effects of reduced insulin secretion and reduced insulin transport into the CNS could contribute to increased energy intake and obesity observed in animals and humans consuming high fat diets.

Multiple physiological systems are in place to maintain energy levels when energy levels fall, nutrients are not available, or during exercise. Moreover, the normal reliance of the control system for energy homeostasis on long-term energy signals including endogenous insulin and leptin is best demonstrated under conditions when their influence is removed (286). Blood

glucose levels are tightly regulated in the range of ~5 mM. When glucose begins to fall, glucagon is secreted from the pancreas, which activates systems designed to utilize stored energy and minimize energy expenditure. Acting on hepatocytes, glucagon activates the enzymes that break down glycogen and release glucose, while also increasing hepatic gluconeogenesis. Importantly, hepatic glucose production ensures a sufficient supply of glucose to the CNS in the fasted state. The exact contribution of glycogenolysis and gluconeogenesis to glucose production remains controversial. Glycogenolysis occurs within 2-6 hours after a meal in humans, and gluconeogenesis has a greater importance upon prolonged fasting.

In adipose tissue, the fall of insulin and rise in glucagon results in decreased lipid storage. Furthermore, this enhances the mobilization of FA's from adipose tissue and as a result the availability of FA's to the liver. The increase in glucagon enhances the oxidation of these FA's by stimulating pathways for fat oxidation inside the liver (Fig. 4).

In cases of prolonged energy deprivation such as fasting, leptin secretion is minimal, reflecting the low metabolic activity of adipocytes. This leptin deficiency causes the brain to sense starvation (despite massive obesity), driving hunger, suppressing energy expenditure, and

51 inhibiting reproductive competence, all advantageous adaptations in the context of starvation. Additionally, when an individual loses weight, less insulin is secreted, consequently increasing food intake until body weight and insulin levels are restored. Finally, glucose transporter levels are also down-regulated specifically in adipose tissue under fasting conditions so as to decrease unwanted clearance of circulating glucose by the adipocytes (304).

Interestingly, leptin possesses both an insulin-like effect on postprandial glycogenolysis and a glucagon-like effect on gluconeogenesis, exerting its effect on the regulation of liver glycogen stores in the fasted as well as the fed state (305). Thus, depending on the hepatic pathways involved, leptin has both insulin-like and insulin-antagonistic effects on glucose metabolism.

The observation that leptin levels are elevated in the vast majority of obese individuals has led to the hypothesis that most obese subjects are resistant to the actions of leptin. Leptin resistance could result from decreased leptin transport into the CNS or to impaired signaling downstream of the leptin receptor (306). Together, these observations are consistent with the hypothesis that the biological impact of leptin is more pronounced when circulating leptin levels are decreasing than when elevated.

During a transition from the resting state to exercise, a profound and rapid shift in whole-body fuel metabolism must occur in response to the rapid increase in the metabolic demands of skeletal muscle. In response to exercise, the secretion of glucagon and insulin from the pancreas generally increase and decrease, respectively. This element of the endocrine response to exercise is critical for the maintenance of glucose homeostasis during exercise. The rise in glucagon and fall in insulin are important for the stimulation of hepatic glycogenolysis and gluconeogenesis. In addition, glucagon also increases hepatic fat oxidation during exercise. The fall in insulin enhances the mobilization of FA's from adipose tissue and makes them more available to the liver, while glucagon enhances the oxidation of these FA's by stimulating fat oxidation inside the liver. Hepatic amino acid extraction and the channeling of amino acid carbons to glucose are also increased. Because of the important roles that glucagon and insulin play, any physiological or pathological condition that affects their secretion or efficacy impacts the metabolic response to exercise.

Taken together, it is evident that several tissues are involved in the buffering of lipid and glucose stores, both in times of energy excess and depletion. Moreover, the brain and periphery carry on a constant conversation; the periphery informing the brain about its metabolic needs and the brain providing for these needs through its control of pathways involved in energy intake, expenditure and storage. Additionally, the cross-talk between peripheral metabolic tissues plays a crucial part in adaptive energy storage and utilization.

C. Hormonal signals

Much like leptin, a large number of factors produced by adipose tissue, as well as other classic endocrine hormones function in concert with the CNS, liver, pancreas, and muscle, in the coordination of energy homeostasis and fuel metabolism. Notably, following the recent identification of several novel hormones implicated in metabolic homeostasis, inter-tissue communication has emerged as a central theme in the study of normal and pathologic fuel

52 Brain Fed Energy expenditure Fasting Hunger

Leptin

Adipose tissue Muscle GLUT4 LPL FFA FAS Glucose uptake HSL

Glucose

Insulin Glucose production Glycogenesis Fatty acid oxidation

Glucagon

Pancreas Liver

Fig 4. Model summarizing integrated metabolic homeostasis under fed and fasted conditions. Adipose tissue, pancreas, liver, muscle, and the brain communicate with each other to regulate energy balance. Following a meal (blue arrows), insulin from the pancreas stimulates glucose uptake and energy storage mechanisms, including increased glycogenesis, as well as lipoprotein lipase (LPL) and fatty acid synthase (FAS) activity. Leptin and insulin levels, which correlate with energy stores, act on the brain to increase energy expenditure and decrease hunger. Under fasting conditions (red arrows), glucagon is released from the pancreas, which decreases glycogenesis and hepatic glucose production for utilization by the brain and other peripheral tissues. Additionally, energy is partitioned away from adipose tissue by decreasing the activities of glucose transporter 4 (GLUT4), LPL, and FAS. Adipocyte lipolysis is increased via the actions of HSL, releasing free fatty acids (FFA) as an energy source for tissues. As leptin and insulin levels drop, the brain decreases energy expenditure, while increasing hunger.

53 physiology. It should be noted that it is nearly impossible to fully cover each factor with respect to metabolism, partially because their contributions are unknown, but also because so many systems are involved.

As mentioned earlier, numerous gastrointestinal hormones have been implicated in food intake regulation. With the exception of ghrelin, other peptides uniformly inhibit food intake (285). These factors include glucagon and amylin, which are released from the endocrine pancreas, as well as enterostatin, somatostatin, gastric inhibitory peptide, gastrin-releasing polypeptide/bombesin, and glucagon-like peptide 1. Many of these peptides and their receptors are also present in regions of the CNS involved in the regulation of feeding behavior. Most gastrointestinal peptides that inhibit feeding exert their effects at much lower doses in the brain than when administered peripherally. It is likely that the peripheral and CNS production and actions of gastrointestinal peptides represent parallel pathways in the modulation of feeding behavior.

Besides factors secreted from the gastrointestinal system, there are other classical endocrine regulators for food intake and energy balance. Glucocorticoids, named for their involvement in glucose metabolism, play an important catabolic role during fasting (307). By stimulating hepatic gluconeogenesis and inhibiting glucose uptake, glucocorticoids increase and maintain normal concentrations of glucose in blood. Moreover, by stimulating lipolysis, glucocorticoids increase the availability of fatty acids for use in production of energy in tissues like muscle, while the glycerol released provides a substrate for gluconeogenesis. Other adrenal-derived hormones, the catecholamines, are also potent stimulators of lipolysis (308). Although glucocorticoids are primarily catabolic in the periphery, they have anabolic effects in the CNS where they increase food intake (309). The changes in hypothalamic neuropeptide systems that inhibit food intake in response to insulin and leptin are for the most part opposed by glucocorticoids (310). Glucocorticoids are believed to interact with insulin and leptin in the long-term regulation of energy intake and body adiposity (311).

Thyroid hormones are essential for normal metabolism (312). Individuals with hyperthyroidism experience increased appetite and food intake, usually with accompanying weight loss. The mechanisms by which thyroid hormones influence feeding behavior are not well understood, however the increase may be mediated by thyroid hormone perturbations of energy expenditure. It has been proposed that by stimulating basal metabolic rate, thyroid hormones create a state of negative energy balance that is associated with loss of body fat and reduced circulating leptin and insulin, which leads to increased energy intake (313). Recent studies indicated that the increase in food intake induced by thyroid hormone is secondary to decreased hypothalamic ATP content (314). In contrast, hypothyroidism decreases basal metabolic rate and leads to weight gain and reduced food intake. However, this does not cause marked obesity, perhaps because weight gain is limited by increased insulin and leptin. In addition, the removal of feedback inhibition of hypothalamic thyrotropin-releasing hormone by the reduced levels of thyroxine occurring in hypothyroidism limits weight gain because central administration of TRH suppresses food intake.

While leptin is proposed to be an adiposity signal to regulate food intake and body weight during health, proinflammatory cytokines such as certain interleukins and TNFα inhibit feeding during

54 acute disease. Feeding inhibition in disease is a component of the acute-phase response characterized by local and systemic reactions, and may have a beneficial effect by decreasing the availability of nutrients to pathogenic organisms. Notably, although cytokines are certainly involved in the anorexia associated with infection and cancer, their role in regulating food intake under physiological conditions (i.e., in the absence of neoplasia or inflammation) is not well understood (315).

Section III. Prolactin

PRL is a peptide hormone synthesized primarily by the pituitary gland. It is classified as a cytokine, which represents a diverse group of molecules, primarily growth factors and hormones, many of which are similarly structured as a four-helix bundle, arranged in an up-up-down-down topology. Cytokines collectively exert a wide range of actions, including immune responses, inflammation, hematopoiesis, oncogenesis, neurogenesis, and metabolism. Members of this family transmit their signal via multi-subunit receptor complexes that are also structurally similar and activate common transduction pathways.

Many individual cytokines are themselves pleiotropic and some have overlapping actions. Pleiotropic actions can be explained by the presence of receptors for a cytokine on multiple cell types or by a cytokine having the ability to activate multiple signaling pathways. Overlapping actions by different cytokines can result from similar cellular distributions of specific receptors, binding to receptors for other ligands, as well as sharing of signaling pathways, which particularly occurs when different receptors share similar motifs that mediate the coupling to the same signaling pathways.

A. Gene structure and regulation of expression

PRL is a member of the gene family that includes growth hormone (GH) and the placental lactogens (PL) (316). Members of this family share similarities in their amino acid sequences, and possess common immunological and biological properties. The genes of these hormones evolved from a single ancestral gene by duplication and sequence divergence (317). A 16- kilobase fish gene coding for somatolactin shares 24% identity with PRL and GH, suggesting that it may be the ancestral precursor for both PRL and GH (318).

In rodents, the PRL gene is approximately 10 kb long and is composed of 5 exons and 4 introns (Fig. 5) (319). Two distinct promoter domains upstream of the rat gene are important for regulation and tissue-specific expression (320). A proximal promoter lies 5’ of the transcription start site, and a more distal enhancer lies further upstream. Highly conserved DNA sequences located in the PRL promoter direct tissue-specific expression of the PRL gene through binding of pituitary transcription factor-1 (Pit-1), a pituitary specific protein containing an N-terminal transactivating domain and a POU homeodomain (321). The human PRL (hPRL) gene, located on chromosome 6, contains an additional exon, 1a, which is located approximately 5.8kb upstream of the coding region (322). Exon 1a is only transcribed in extrapituitary sites, where it is spliced to exon 1b, producing a transcript that is 134 bp larger in the 5’ untranslated region (323). Extrapituitary transcription of PRL is regulated by the superdistal promoter, which lies

55 upstream of exon 1a (324). Pituitary and extrapituitary derived PRL are identical, as judged by biochemical, biological and immunological criteria.

Numerous hormones, neurotransmitters, growth factors, and steroids affect PRL expression. Acting either directly on response elements in the promoter/enhancer region (e.g. estradiol), or via intracellular signaling pathways that target the promoter (e.g. TRH), they mediate transcription of the gene. For more information on the regulation of pituitary, as well as extrapituitary PRL expression, the reader is directed to reviews by Gourdji and Ben-Jonathan (320;325).

B. Protein structure and variants

PRL typically exists at approximately 23-kDa molecular weight and 197-199 amino acids, with variable sequence homology among species (326). Structurally, PRL is a single-chain polypeptide containing six conserved cysteine residues that form three disulphide bridges that form two short loops at the N- and C-terminals and a large intermolecular loop (Fig. 5) (327). Using three-dimensional and four-dimensional NMR spectroscopy, Keeler et al. have shown that PRL adopts the predicted “up-up-down-down” four-helical bundle topology, similar to GH and other hematopoietic cytokines (328). Post-translational modifications including glycosylation, phosphorylation, and proteolytic cleavage, which generates several variants. Glycosylated PRL, with lower biological activity than non-glycosylated PRL, has been detected in several species, including mammalian and avian (329). Varying from 1% to more than 50%, the extent of glycosylation differs among species and may also depend on reproductive state (330). Phosphorylated isoforms of PRL are secreted by pituitary cells in vitro, and have been detected in bovine and murine pituitary glands. However, it is not known if phosphorylated PRL is secreted into the plasma in vivo (331;332). Receptor binding is decreased by phosphorylation, resulting in dramatically lower biological activity than non-phosphorylated PRL (333). Interestingly, recent in vitro and in vivo studies have demonstrated potent anti-angiogenic and anti-tumorogenic effects of a molecular mimic of phosphorylated PRL (334). Proteolytic cleavage of 23 kDa PRL can produce many isoforms including the N-terminal 22-, 16-, and 14 kDa variants, which have been extensively investigated (327). The 16 kDa isoform has received most attention, and is proposed by Clapp et al. to be the main product of proteolysis by target tissues (335). Its estimated mitogenic and lactogenic potencies were 65% and 10% those of native PRL, respectively, and it reportedly acts as an anti-angiogenic factor, inhibiting endothelial cell proliferation and capillary formation (336). Recent studies have shown that 16 kDa PRL can act directly on endothelial cells to inhibit several other processes associated with angiogenesis, including cell-cell and cell-matrix interactions, and the degradation of extracellular matrix. Additionally, the ability of 16 kDa to inhibit angiogenesis results from its ability to promote endothelial cell apoptosis (337). The proportions of PRL variants vary with physiologic, pathologic, and hormonal stimulation.

56 Extrapituitary start site Pituitary start site

-10kb -6 0 +10kb

5’- 1a 1b 2 3 4 5 3’-

Pituitary

Extrapituitary

N Exon Signal peptide C PRL Promoter PRL transcript Protein Untranslated region

Figure 5: Diagram of the PRL gene, PRL mRNA transcript and mature PRL protein. Arrows designate transcriptional start sites for proximal pituitary promoter and the superdistal extrapituitary promoter. Transcription from either promoter produces mRNAs with identical protein coding sequences but differing in the 5’ untranslated sequences (5’UTR). The mature PRL protein shows the four helix bundle configuration.

57 C. Pituitary lactotrophs

The best-recognized site of PRL production is the lactotrophs of the anterior pituitary (AP) gland. These cells were originally described by Herlant et al. in rat pituitaries in 1964 (338). Their presence was later confirmed in the AP of mice and humans using species-specific PRL antibodies (339;340).

Lactotrophs constitute a heterogeneous cell population, differing in morphology, distribution, function, and responsiveness to stimuli. They are derived from a cell lineage that includes somatotropes and thyrotropes, which appears during middle to late gestation. The development of specific cell types in the pituitary is directed by transient signaling gradients that induce nuclear mediators of cell type commitment, including transcription factors acting as repressors or activators, and their associated coregulators. Anterior pituitary (AP) cell types are initially positionally determined, with the somatotrope/lactotrope cells arising caudomedially and gonadotropes more rostroventrally, corticotropes ventrally, and melanotropes dorsally (341). For each cell type to progress beyond initial patterning by transient signaling gradients, induction of additional specific transcription factors is required. These transcription factors include Pit-1 for somatotropes, lactotropes, and thyrotropes; the orphan nuclear receptor SF1 and Egr-1 for gonadotropes; and T-pit and possibly Stat3 for POMC gene-expressing cells (342). Pit-1, a POU domain protein directly controls regulatory genes, including receptors involved in growth and homeostatic control (343). The Pit-1 lineage can be converted to alternative fates before E17.5 but exhibits a permanent cell-autonomous commitment after E17.5, when Pit-1 gene regulation to a Pit-1-auto-regulated enhancer (341). Lactotrophs are the last cell type to differentiate, and at day 5, comprise only 5% of the total AP cell population, but increase significantly in number between postnatal days 6-20 (344). While there are no differences in lactotroph cell population between young males and females, sexual differences develop around puberty, and are dependent on reproductive status. In adult females at proestrous, about 50% of AP cells are lactotrophs, compared to 35% in adult males (344).

In examining lactotroph morphology, De Paul et al. noted cells of various sizes, as well as shapes, ranging from polyhedral, to angular, round, or oval (345). The distribution of lactotrophs within the AP is uneven, with the highest density found adjacent to the intermediate lobe, and fewer cells located in the lateroventral portion of the AP (346). Interestingly, Boockfor et al. demonstrated that lactotrophs from specific regions of the AP respond differently to certain stimuli (347). Density gradient separation of AP cells identified two lactotroph subpopulations that differed in their resting membrane potential, as well as basal and stimulated rates of PRL secretion (348). Although they are considered endocrine cells, lactotrophs also exhibit several traits characteristic of neurons, including spontaneous electrical activity and stimulus-induced current influx (349).

A minority cell type found in the anterior pituitary, the mammosomatotrophs produces both GH and PRL (350). As suggested by Frawley, these dual secreting cells may possess the ability to differentiate into lactotrophs or somatotrophs (351). Additionally, Boockfor et al. observed that estrogen can convert somatotrophs into mammosomatotrophs, indicating that these cells may indeed interconvert between single- and dual-secreters (352).

58 While the pituitary is the primary site of PRL production in rodents, numerous extrapituitary production sites exist in humans. PRL has been detected in the mammary gland, decidua, myometrium, and other tissues in humans (325). Within the human brain, PRL has been detected by immunohistochemistry in the hypothalamus, cerebral cortex, hippocampus, amygdala, septum, cerebellum, and choroid plexus. It is found in fluid compartments, including cerebrospinal fluid (CSF), amniotic fluid, tears, and milk, originating from both pituitary and extrapituitary production sites. Interestingly, abdominal adipose tissue has recently been reported to produce PRL and express the PRLR in humans (353;354). The wide distribution of both PRL and its receptor suggest that in humans, PRL acts in an autocrine/paracrine manner. The placenta produces an assortment of PRL-like molecules called placental lactogens (PL) that are structurally similar to PRL. PL binds to the PRLR and alters maternal carbohydrate and lipid metabolism to ensure the optimal supply of nutrients to the fetus and utilization of the nutrients by fetal tissues (355).

Several lactotroph cell lines have been cloned from pituitary tumors and are widely used as in vitro models to study the synthesis, processing, and secretion of PRL. The most extensively studied cell line, GH, is a mammosomatotroph isolated from a radiation-induced pituitary tumor in a Wistar-Furth fat (356). Several subclones have been produced, including the GH1, GH3 and GH4 cell lines that produce both PRL and GH in different ratios. GH cells continuously synthesize and secrete PRL due to limited storage capacity. Although DA is the primary inhibitor of PRL release in vivo, GH cells are unresponsive to DA agonists due to the lack of a functional dopamine-2 receptor (D2R). The MMQ cell line, also commonly used to study the regulation of PRL synthesis and secretion was established from the estrogen-induced rat pituitary tumor 7315a, and exclusively produces PRL (357). MMQ cells are nonadherent, express a functional D2R, and their PRL secretion is inhibited by DA or DA agonists. PR1 is an estrogen-responsive cell line derived from diethylstilbestrol-treated F344 female rats that shows a unique response to estrogen. The concentration of estrogen required for PR1 proliferation is 1000-fold lower than that required to increase expression of the PRL gene (358). With no human PRL-producing cell line in existence, investigating and understanding the mechanisms that regulate PRL expression and secretion in humans is difficult.

D. Regulation of pituitary PRL secretion

PRL secretion is controlled by a large variety of stimuli provided by the environment and the internal setting. Important regulators of PRL secretions include suckling, stress, ovarian steroids, and many others. These stimuli are received by the hypothalamus, which coordinates an appropriate secretory response.

The neuroendocrine control of PRL secretion is a multifactoral process that involves both stimulatory and inhibitory molecules. Pituitary lactotrophs are active secretory cells that maintain a high rate of basal secretion in the absence of an inhibitory signal. Lesions of the hypothalamus or ME, or disconnection of the pituitary gland from the medial basal hypothalamus augment PRL release (359;360). Similar effects on PRL release are observed in vitro from isolated pituitary fragments or dissociated AP cells, indicating that a factor(s) released from the hypothalamus inhibits PRL secretion. This is unique to PRL since the release of all other major pituitary hormones is regulated by stimulation.

59

Pituitary PRL is primarily subjected to tonic inhibition by dopamine (DA) (361). DA is a neurotransmitter/neurohormone that participates in the regulation of numerous functions, including movement, reward, and PRL secretion. It is produced primarily in the striatum, but is also produced in the arcuate and periventricular nuclei of the hypothalamus. Physiologically a low basal rate of PRL secretion is primarily maintained by DA release from hypothalamic tuberoinfundibular (TIDA) neurons (362). These cells project from the hypothalamus, where their perikarya are located, to the external zone of the ME, where DA is released. DA is delivered from the ME into the sinusoid capillaries of the anterior pituitary through long portal vessels, where it binds to the D2R on lactotrophs.

The rate of DA biosynthesis and its release from TIDA neurons are well-known in the rat based on measurements of the activity of tyrosine hydroxylase, the rate-limiting enzyme in DA biosynthesis, the turnover rate of DA, and the concentration of 3,4-dihydroxyphenylacetic acid (DOPAC), a DA metabolite. Alterations in the activity of TIDA neurons reflect changes in PRL release and an inverse relationship between the concentration of DA in portal blood and plasma PRL is well documented (361).

PRL regulates its own secretion through a short negative feedback loop. TIDA neurons, expressing high levels of the PRLR, are activated by PRL which bypasses the blood-brain barrier and enters the CSF through the choroid plexus. In this feedback system, elevated serum PRL increases hypothalamic DA release, thus increasing the inhibitory signal and decreasing pituitary PRL secretion. Tyrosine hydroxylase (TH), an enzyme involved in DA synthesis, is one of the targets of PRL action within the hypothalamus. Hyperprolactinemia and hypoprolactinemia increase and decrease, respectively, TH expression in TIDA neurons and DOPAC accumulation in the ME (363). In female rats, long-term estrogen treatment decreases TIDA responsiveness to PRL. Central PRL administration can increase DOPAC accumulation in the ME, while PRL antibodies are effective in neutralizing basal plasma PRL, as well as decreasing TIDA activity and DOPAC buildup in the ME.

Besides the TIDA neurons, the ME contains two other major dopaminergic systems: THDA (tuberohypophyseal), and PHDA (periventricular hypophyseal) neurons. These neurons pass through the internal zone of the ME and infundibular stalk and innervate the IL and NL, respectively. DA released in the NL from THDA and PHDA neurons reach the AP via the short portal vessels and contribute to PRL inhibition (364). Indeed, neural lobectomy in rats results in a rapid elevation in PRL that is reversible by DA injection (365). Furthermore, during lactation and suckling, DA content and turnover rate in the ME and NL decreases. Additionally, the NL maintains PRL inhibition despite the separation of the AP from the hypothalamus resulting in the loss of TIDA input. In sheep, the NL may contribute most of the DA, via the short portal vessel that is responsible for the regulation of PRL. Thus, alterations in hypothalamic or NL DA levels may result in differential regulation of PRL release in some species.

Although hypothalamic DA is the predominant inhibitory molecule regulating PRL secretion, factors produced within the pituitary gland can also decrease PRL production and release. TGFβ1, a member of the TGFβ family, inhibits plasma PRL and pituitary weight following intrapituitary administration (366). In primary AP cultures, TGFβ1 decreases basal and estradiol-

60 induced PRL production and release. Additionally, TGFβ1 reduces basal and calcium-stimulated PRL mRNA levels in GH3 cells via a Pit-1 independent mechanism.

Other inhibitors of PRL secretion include calcitonin, endothelin and gamma aminobutyric acid (GABA). One report showed that calcitonin inhibits basal and TRH-induced PRL mRNA and release from rat AP cells by inhibiting a calcium-inositol tri-phosphate signaling pathway (367). More recent studies show that i.c.v. administration of calcitonin increased TH activity of TIDA neurons in the ME, while completely suppressing the PRL response to suckling (368). Endothelins have been identified in the pituitary and inhibit PRL release from AP cells, although the underlying mechanism is unknown. GABA regulates PRL secretion by either altering the dopaminergic tone or directly acting on AP cells. GABA inhibits PRL secretion by acting on GABA-A and GABA-B receptors on AP cells, while it PRL secretion is enhanced via direct stimulation of the GABA-C receptor (369). However, the participation of the cholinergic system in the regulation of PRL release is controversial, and the observed effects vary with the site of administration, the dose, and physiological conditions.

Although dopamine is recognized as the primary inhibitor of PRL secretion, its precise mechanism of action is still not clear. Based on observations in vitro, DA exerts its effects on lactotrophs in several ways. DA treatment rapidly induces membrane hyperpolarization, causing inactivation of voltage-gated Ca2+ channels and increased voltage-dependent K+ currents. This decreases the excitability of the cell and inhibits PRL release from secretory granules, as demonstrated by Lledo et al. (370). Minutes to hours later, DA suppresses adenylyl cyclase activity, resulting in decreased cAMP levels and altered PRL gene expression (371). Finally, within 24 hours of DA treatment, proliferation decreases and cell morphology is altered (372).

Many factors can elevate PRL expression/secretion. Unlike other AP hormones, a singular hypothalamic PRL releasing factor (PRF) has not been identified. Some compounds affect secretion by altering dopaminergic regulation of PRL, while others directly affect PRL expression/secretion. The list of factors is long; therefore I will only mention a few of the most- well characterized secretogogues. TRH, originally isolated as a hypophysiotrophic factor that stimulates thyroid-stimulating hormone (TSH) secretion from pituitary cells, is an effective stimulator of PRL release both in vitro and in vivo (373;374). TRH neurons from the PVN innervate both the ME and posterior pituitary (PP), and TRH levels in portal blood show some correlation to elevated PRL during the afternoon proestrous and suckling. Moreover, intravenous administration of TRH results in a rapid elevation in serum PRL (375). The action of TRH on PRL release occurs through activation of a G-protein coupled receptor, which leads to increased intracellular calcium and subsequent release of PRL from storage secretory granules (376).

Vasoactive intestinal peptide (VIP), a potent vasodilator produced in numerous tissues, including the suprachiasmatic nucleus, also increases PRL release (377). Acting directly on VIP receptors on lactotrophs, it up-regulates PRL gene expression by increasing cAMP levels and activating PKA. VIP is less potent than TRH and the temporal stimulation of PRL is delayed and more gradual. VIP neurons have been identified in the PVN and their activation by serotonergic neurons from the dorsal raphe are thought to mediate stress-induced PRL release (378). The effect of TRH and VIP on PRL release is additive, indicative of the independent signaling

61 pathways activated by these peptides, with VIP exerting its effects via cAMP-mediated activation of PKA.

Oxytocin is a nonapeptide that is released during physiological states during which PRL is also elevated, including labor, nursing, and orgasm. Synthesized in the PVN and supraoptic nucleus, it can reach the anterior lobe from the ME via the long portal vessels, or from the neural lobe (NL) by short portal vessels (379). However, it is still uncertain if the synchronized release of PRL and oxytocin are concomitant. A direct action of oxytocin on PRL release supported by in vitro studies, but the relative weak potency of oxytocin compared to other secretogogues makes its contribution to the physiological regulation of PRL questionable. For a more in depth discussion on the known regulators of PRL secretion, the reader is directed to the review by Freeman (380).

Recently, another PRF was discovered when PP lobectomized rats were used to investigate the role of the THDA neurons in the suckling-induced PRL rise. Basal PRL release was elevated in lobectomized lactating rats, confirming that dopamine from the THDA neurons contributes to the tonic inhibition of PRL release. However, suckling, with or without oxytocin administration, failed to increase plasma PRL levels, indicating the presence of a PRF in the PP. Additional studies have shown that the activity of this PRF is localized in the IL but not in the NL, hypothalamus or cerebral cortex. PP extracts stimulated PRL release in a rapid, dose-dependent and hormone-specific manner both in vivo and in vitro, while co-culturing posterior pituitary cells and lactotrophs resulted in a dramatic increase in PRL mRNA levels. Unfortunately, attempts to isolate and identify this PRF have been unsuccessful.

E. Physiology of PRL

PRL produced in the pituitary acts as a classical hormone. It is secreted by the gland into the circulation, transported by the circulatory system, and acts on peripheral tissues. The patterns of PRL secretion are species-specific, exhibit sexual dimorphism, and vary under different physiological conditions and in response to numerous stimuli. Additionally, PRL secretion involves complex interactions of regulatory factors and plasticity of neuronal responsiveness.

The best-known physiological stimulus for PRL release is suckling applied by nursing during lactation. The suckling response is a widely studied reflex, comprising a neural afferent and a humoral efferent component. The neural afferent component is initiated by the stimulation of numerous mechanoreceptors in the nipple and surrounding areas of the skin during suckling. It is assumed that the signal is mediated through the spinal cord and brain stem up to the hypothalamus where information is transformed into a humoral efferent reflex answer. The latter is expressed via changes in the secretion of PRL releasing and/or inhibiting factors, including DA, TRH, VIP, oxytocin, and others produced by hypothalamic neurons. In rodents, PRL concentrations increase and the level of DA in the ME is reduced within minutes of the initiation of nursing. PRL levels peak within 10 minutes, and remain elevated at a constant level throughout the suckling (381). The quantity of PRL released correlates with the intensity of the suckling stimulus. Pup separation decreases plasma PRL and increases the concentration of DA in the ME and AP. The concentration of DA continues to rise in these regions as the time interval

62 of pup separation and non-suckling increases (382). When nursing is completed, PRL levels rapidly fall in proportion to the metabolic clearance rate of the hormone (383).

Studies have implicated serotonin in the suckling response, since there is a rapid decrease in the hypothalamic concentration of serotonin and an elevation of its metabolite, 5-hydroxyindolacetic acid, simultaneously with the release of PRL (384). Additionally, selective lesion of the PVN markedly reduces the PRL response to serotonin. Other reports have shown that blockade of the glutamate receptors causes a decline in plasma PRL concentration of suckling mothers, leading to the conclusion that the endogenous glutamatergic system has an important role in suckling- induced PRL elevation and in the maintenance of high PRL levels during lactation (385).

PRL levels fluctuate during the reproductive cycle. In rodents, PRL secretion remains low during most of the estrous cycle but surges along with luteinizing hormone (LH) during the afternoon of proestrous, via an estrogen-mediated mechanism (386). While the LH surge is symmetrical, the PRL surge is not, displaying a fast, sharp peak followed by a prolonged plateau phase that gradually declines (387). During the late afternoon and evening of proestrus, the short-term inhibitory effect of progesterone on TH activity in TIDA neurons prolongs the preovulatory PRL surge via a dopaminergic mechanism (388).

In the absence of mating, the lack of PRL needed to maintain the corpus luteum results in its disintegration. However, if mating does occur, or if appropriate stimulus is applied to the uterine cervix, the corpus luteum is rescued by twice daily PRL surges, whose timing is maintained by the suprachiasmatic nucleus (389). During pseudopregnancy, they end after 12 days, followed by resumption of the estrous cycle. If pregnancy occurs, these PRL surges end after 10 days due to negative feedback on the AP from rising levels of PL. PRL remains low during mid- to late- gestation, until a surge during the dark period just before parturition. In early lactation a PRL- induced increase in TH activity leads to negative feedback, but this effect is lost by mid- lactation. The negative feedback during early lactation is overridden by the inhibitory effect of suckling on dopaminergic activity (390).

The physiology of PRL in humans is different from that in rodents. Circulating PRL levels remain fairly constant during the menstrual cycle (391). There is no preovulatory surge, and estrogen has little effects on PRL release. Maternal PRL levels rise gradually towards term, reaching levels 10 fold higher than in non-pregnant women (392). Elevated PRL during pregnancy is a functional reflection of lactotroph hypertrophy and hyperplasia, which is induced by estrogen (393). The rise in PRL is necessary for preparation of the breast for lactation, which is inhibited by the high levels of progesterone maintained during pregnancy. During mid- pregnancy, progesterone stimulates the production of large amounts of PRL from the decidua, which is secreted into the amniotic fluid (394).

Human fetal PRL levels are low during early gestation, rising gradually and reaching maximal levels before birth (395). Another rise takes place birth and persists for several days. Similar to rodents, there are no sex differences in serum PRL levels prior to puberty, but female PRL levels are twice as high as males after puberty (396). Suckling is also the most potent stimulus for PRL release in humans (397). Physiological stimuli can also increase circulating PRL levels. Stressors such as surgery, anesthesia, and exercise increase PRL release in men and women. Other

63 external stimuli, including odor and sound affect PRL production. There is a nocturnal rise in PRL that is regulated by sleep rather than by light or circadian rhythm (398).

Section V. Prolactin Receptors

A. Protein Structure

The PRLR belongs to the class 1 cytokine receptor family, which also includes receptors for GH, leptin, interleukins, and several other factors (399). Although members of this family are genetically unrelated and show variable degrees of sequence homology, they share functional and structural properties (400). Each receptor is a single-pass transmembrane protein containing an extracellular ligand-binding domain, a transmembrane domain, and an intracellular domain that mediates signaling (Fig. 6) (401).

The extracellular domain of the PRLR is ~200 amino acids and shares considerable sequence homology with other cytokine receptors. It contains two subdomains, an NH2-terminal region referred to as D1, and a membrane-proximal portion called D2 (399). Both subdomains exhibit similarities with type 3 fibronectin, which directs the ligand-receptor interactions in cytokine receptors. D1 contains two pairs of disulphide bonds (between Cys12- Cys22 and Cys51- Cys62) and D2 contains a “WS-motif” (Trp-Ser-x-Trp-Ser). The disulphide bonds and the WS motif are crucial for proper folding and trafficking of the receptor (401). Three glycosylation sites exist in the extracellular region of the PRLR, two of which are in the D1 domain that are also necessary for proper cell surface targeting (402). The transmembrane domain, 24 amino acids in length, is well conserved among species, but it is not known whether this region of the PRLR is involved in receptor function (401).

Box 1 within the membrane proximal cytoplasmic region is a proline-rich hydrophobic motif. This region contains a Pro-x-Pro sequence that adopts the typical folding of SH3-binding domains, and is recognized by signal transducers during receptor signaling (403;404). The last proline in the Box 1 segment is reportedly critical for JAK2 association and subsequent signaling (405). Box 2 lies further from the membrane and consists of a succession of hydrophobic, negatively charged, and then positively charged residues. It is less conserved than Box 1 and is not present in the short form of the receptor (401).

While the extracellular and transmembrane domains of all PRLR are primarily the same, receptor isoforms differ considerably in the intracellular domain. These isoforms are a result of alternative splicing and proteolysis. In mice, one long (L) and three short (S1, S2, S3) forms have been discovered, the short forms only differing by a few amino acids in the C-terminal of their cytoplasmic tail (406). Three isoforms have been described in rats, including the short, intermediate, and long forms, while in humans a long (L), an intermediate (I) and two short isoforms (S1a and S1b) have been detected (407-411). In addition to the membrane-bound PRLR, soluble isoforms which contain only the extracellular domain have been identified (412;413). In several species, soluble PRLR functions as a PRL binding protein, reportedly engaged with 36% of serum PRL (414;415).

64 Long S-1 S-2 S-3

D1 D1 D1 D1

D2 D2 D2 D2

Box 1 Box 1 Box 1 Box 1

Disulphide Bond WS motif Cell membrane Unique sequence

Figure 6: The PRLR is a single pass transmembrane protein belonging to the class I cytokine receptor subfamily. The mouse PRLR exists as four isoforms: long (L), and three short isoforms (S-1, S-2, and S-3). The extracellular domain consists of 210 amino acids and contains two fibronectin like regions, D1 and D2. The D1 domain contains two N-linked glycosylation sites and two disulphide bridges, while the D2 domain has a conserved WS motif (tryptophan-serine-X-tryptophan-serine). The intracellular domain varies among the isoforms. The membrane proximal region contains a proline rich hydrophobic motif known as Box 1 that adopts the typical folding of SH3 binding domains.

65 B. Gene structure

The gene encoding the PRLR contains a common noncoding exon corresponding to the 5'- untranslated region (UTR) which precedes the translation initiation codon. It is followed by 11 coding exons, the last four of which are alternatively spliced to produce the four isoforms of the receptor. The cytoplasmic region of the receptor is encoded by exon 10 for the L-PRLR, exon 12 for PRLR-S1, exon 11 for PRLR-S2, and exon 13 for PRLR-S3 (416).

Regulation of PRLR transcription is achieved via several tissue-specific promoters that code for distinct first exons (417;418). In rats, three gene promoters (PI, PII, and PIII) were identified that regulate tissue-specific transcription of three distinct first exons, E11, E12, and E13. Promoter I is specific for the gonads, promoter II for the liver, and promoter III is common to several tissues. PI is localized to the 152-base pair 5' of the transcriptional start site at -549. It contains a steroidogenic factor 1 (SF-1) element that binds the SF-1 protein to activate the promoter (419). A CCAAT box contributes minimally to basal activity in the absence of the SF-1 element, and two adjacent TATA-like sequences act as inhibitory elements. Deletion analysis revealed a binding site for hepatocyte nuclear factor 4, a liver-enriched transcription factor that activates PII (417). A C/EBPβ binding domain and an Sp1 element within the promoter contribute individually to PIII activation in gonadal and non-gonadal cells. The wide expression of these two transcription factors may explain the common utilization of PIII in multiple tissues and in different species (420).

Five promoters have been identified in the mouse, the first three of which were detected by genomic DNA cloning. An additional two promoter sequences were identified using rapid amplification of cDNA ends-PCR (RACE-PCR) (416). The rat and mouse PIII share similar structure and function, however, the mouse PI lacks the functional SF-1 element and hence is inactive. Human PRLR shares one of its promoters/exons (hE13/PIII) with rodents (421). Five additional exons have been identified, hEN1-hEN5, which code for hPN1-hPN5 (422). The hPIII contains identical Sp1 and C/EBPβ elements as in the rodent promoters, shares 81% similarity in rodents (423). hPN1 activity is mediated by domains containing an Ets element and an NR half- site (422).

C. Activation and signal transduction

The extracellular domain of the PRLR binds to one of two sites on PRL. One site involves helices 1 and 4, while another is made from helices 1 and 3 (424). Stoichiometric analysis by Gertler et al. revealed that PRL binds to its receptor to form a 1:2 complex, which is necessary for activation of the receptor (425). The initial binding site first binds to one PRLR, forming a dimeric complex, then recruits a second receptor to form the active trimeric complex. Gertler demonstrated that the 1:2 complex is unstable, and undergoes rapid dissociation to a 1:1 complex (426).

The activated PRLR signals through several intracellular pathways, the primary and best understood being the Jak2/Stat pathway (Fig. 7). Activated PRLR phosphorylates several cellular proteins, as well as the receptor itself, despite being devoid of any tyrosine kinase activity (427). This activity is mediated via kinases that are constitutively associated with the receptor. For

66 instance, members of the family of Janus Kinases (Jak), including Jak1, Jak2, Jak3, and Tyk2, associate with cytokine receptors to mediate ligand induced signals within the cell (428). The signal transducer and activators of transcription (Stat) are a family of proteins known as major transducers of cytokine signaling (429). Four of the Stat proteins, particularly Stat5a and Stat5b, and to a lesser extent Stat1 and Stat3 are involved in PRLR signaling.

Box 1 in the membrane proximal region of the PRLR intracellular domain is associated with Jak2, which becomes phosphorylated within one minute of PRL binding (430). Activation of Jak2 occurs by transphosphorylation after two Jak2 molecules are brought together by receptor dimerization. When activated, Jak2 phosphorylates tyrosine residues on different target proteins, including the Stat proteins, as well as the PRLR itself. Tyrosine residues on the short form of the PRLR, which does not contain Box 2, are not phosphorylated by Jak2. Stat proteins contain several conserved features, including an SH2 binding domain, an SH3 domain, and a DNA binding domain. The SH2 domain interacts with phosphotyrosine residues on the activated PRLR, as well as with the associated Jak2 proteins. The ability of the short form of the PRLR to signal through Jak2 is debated. While one report indicates that both long and short forms of the receptor associate with Jak2 and activate Stat1, others report the opposite, and suggest that it acts as a dominant negative by heterodimerizing with the long form of the receptor (403;431).

While docked at the PRLR/Jak2 complex, Stat proteins are phosphorylated by the receptor associated Jak kinase. Upon phosphorylation Stat proteins disengage from the receptor and hetero- or homodimerize with another Stat molecule via interactions between their phosphotyrosine residues and SH2 domains. Dimerized Stats translocate to the nucleus where they activate Stat binding motifs, known as gamma-activated sequences (GAS), which are present in the promoters of numerous target genes (432). Observations in Stat knockout mice highlight the importance of the Stats in PRLR signaling. Stat5a knockout mice develop normally, but mammary lobuloalveolar outgrowth during pregnancy is curtailed, resulting in lactational failure due to lack of terminal differentiation of the mammary gland (433). These results demonstrate that Stat5a is the principal and an obligate mediator of mammopoietic and lactogenic signaling. Disruption of the Stat5b gene leads to the major loss of multiple responses associated with the sexually dimorphic pattern of pituitary growth hormone secretion (434). Furthermore, Stat5a/b knockout mice have a defect in the development of functional corporal lutea in the ovary that results in female infertility (435). Female infertility is not observed in the Stat5a or Stat5b -/- mice, demonstrating the functional redundancy of the Stat5 proteins.

Although the Jak/Stat cascade is the primary pathway used by the PRLR, other signal transduction pathways are also activated. For instance, PRL also activates the MAPK cascade, which is involved in the activation of numerous transcription factors/ immediate early genes by phosphorylation (436). Adapter proteins (Shc/Grb/SOS) link the Ras/Raf/MAPK cascade to the PRLR by binding to phosphotyrosine residues of the activated receptor, indicating that the Jak2/Stat and MAPK pathways are interconnected rather than independent (401). Additionally, PRL induces activation of members of the Src family of kinases (SFK), Src, and Fyn (411;437). Stimulation of the Src pathway has been reported to be independent of Jak2 activation and receptor phosphorylation (438). In addition, recent studies have demonstrated c-Src-mediated activation of Fak/Erk1/2 and PI3K pathways (439). Fyn, which is associated with the PRLR, is activated by PRL in certain cell types, and reportedly modulates K+ currents and is proposed to

67 PRL PRL

P P Stat1 Jak2 Jak2 P ? P Stat1 P Jak2 Jak2 P Y Y FYN Y FYN P Stat3 P Y Y P SHC SOS Stat1 GRB RAS P Stat5 P Y Y c-Src RAF K+ Stat3 Stat5 P IRS MEK1/2 PI3K/Akt FAK P Stat 5a/b P PI3K/Akt ERK1/2

P Stat 5a/b P

GAS

Figure 7: Schematic diagram of PRLR signaling pathways. Long and short forms of the mouse PRLR are represented. Jak2 binds to box 1 within the intracellular domain of the receptor. Binding of PRL to the receptor induces receptor dimerization and phosphorylation of Jak2, which results in recruitment of mainly Stat5, but also Stat1 and Stat3. Following phosphorylation and dimerization, Stat5a/b translocates to the nucleus and alters transcription of PRL-responsive genes via binding to the gamma- activated sequence (GAS) motif. Whether the short isoform activates the Stat pathway is unknown. The Jak2/Stat5 pathway is considered the main pathway, but phosphatidylinositol-3-kinase (PI3K)/Akt, and mitogen-activated protein kinase (MAPK) pathways are also stimulated by ligand binding to the PRLR. The MAPK pathway involves the SHC/GRB/SOS/Ras/Raf cascade and is presumably activated by all PRLR isoforms.

68 activate PI3K (437;440). Finally, PRL has also been shown stimulate the IRS/PI3K pathways (441). In all likelihood, in tissues other than the mammary gland, PRL utilizes multiple signaling pathways, the predominance of which is tissue-dependent.

Cytokine-induced signaling pathways such as Jak2/Stat are negatively regulated with respect to both duration and intensity. At least three different classes of regulators are known to contribute to cytokine inhibition, including the protein inhibitors of Stats (PIAS), the Src-homology (SH2)- containing protein tyrosine phosphotases (SHPs), and the SOCS (442). PIAS prevent Stat/DNA interactions by inhibiting Stat dimerization, while SHPs reduce the catalytic activity by causing the dephosphorylation of tyrosine residues critical for receptor signaling (443). The SOCS family consists of eight proteins: SOCS1-SOCS7 and CIS, each of which contains a central SH2 domain and a C-terminal SOCS box (442). Though the function of all SOCS proteins is not completely clear, they primarily block cytokine signaling by steric hindrance of binding sites on the activated PRLR complex that are used to recruit and phosphorylate Stats, particularly Stat5 (444).

The expression of CIS, SOCS1, SOCS2 and SOCS3 is rapidly induced in response to stimulation by a wide variety of cytokines, including PRL. To that end, numerous reports indicate that SOCS proteins inhibit the PRLR/Jak/Stat signaling cascade. SOCS-1 inhibits signaling by binding to Jak2, while SOCS-3 reportedly binds to the PRLR itself to prevent pathway activation (445). Interestingly, SOCS-2 has been shown to antagonize the other SOCS proteins, increasing activated signal intensity (445;446). In vivo studies using specific SOCS knockouts have revealed their physiological importance. SOCS-1 knockout mice exhibit stunted growth and die at 3 weeks of age with lymphopenia, activation of peripheral T cells, fatty degeneration and necrosis of the liver, and macrophage infiltration of major organs (447). SOCS-2 -/- mice grow 30-40% larger than their littermates, while SOCS3 -/- mice die during the embryonic stage of development (448;449).

D. Receptor distribution and regulation of expression

PRL-binding sites or receptors have been identified in a large number of cells and tissues, both in adult and during fetal development (401). The expression of PRLR isoforms vary as a function of the stage of the estrous cycle, pregnancy, and lactation. PRL receptors are distributed in a sexually dimorphic manner in some tissues as well as within the brain. Notably, the PRLR is present in several key tissues regulating metabolism, including the liver, pancreas, and brain. The PRLR was only recently detected in white adipose tissue, and its functional significance in this tissue remains relatively unknown (450).

Hormonal influences on PRLR expression depends on the target organ. Djiane et al. observed that PRLR expression steadily increases in the mammary gland during pregnancy and rises significantly at parturition (451). Studies by Hayden et al. confirmed these results, also reporting that receptor levels declined after the litters were weaned (452). More recently, Camarillo et al. noted increased PRLR levels in the epithelial compartment of the mammary gland during pregnancy and lactation, with no changes within the stromal compartment (453). PRL is able to up- and down-regulate the expression of its own receptor, therefore, the rise in PRLR is attributed to increasing levels of circulating PRL. However the increase in PRLR is limited by

69 high levels of progesterone, which down-regulates receptor expression (454). Thus, following birth when progesterone levels drop, PRL is able to significantly up-regulates its own receptor. PRLR expression patterns in the liver are quite different. Hayden et al. observed that PRLR levels were lower during lactation than during pregnancy or after weaning (452). They suggested that expression of the PRLR is mediated by tissue-specific factors. PRLR expression is detectable in many hypothalamic regions, including the arcuate, periventricular, and medial preoptic nuclei of the hypothalamus (455). Both short and long PRLRs are present in these nuclei, but the expression levels of the two isoforms vary between nuclei (455;456). The choroid plexus contains the highest levels of PRLR mRNA expression and immunoreactivity in the brain, and it has been proposed that PRL binding sites in the choroid plexus are involved in the transport of PRL from the blood to the CSF (457).

PRLR expression is also mediated by steroids. In 1979, Hayden et al. reported that estrogen treatment increases PRL binding in the mammary gland of virgin rats (452). More recent studies support this data, and estrogen appears to be a common inducer of PRLR expression in various types of tissue, including liver, kidney, endometrium, and human breast cancer cells (458). The effects of progesterone vary; as it induces PRL-R mRNA in human endometrial stromal and T47D cells, but inhibits expression in the mammary gland of rats and mice. Bridges et al. have reported that progesterone decreases PRLR expression in the MPOA of the brain, while estrogen alone has no effect (459). However, several other studies have reported higher PRLR expression in the brain of ovariectomized plus estrogen-treated female rats compared to ovariectomized rats only (460).

Sakai et al. first reported that glucocorticoids also regulate PRL binding in the mammary gland. He observed that adrenal ablation caused a reduction in the number of PRLR, an effect reversed by hydrocortisone treatment (461). In agreeance with these observations, infusion of cortisol increases hepatic PRLR in sheep (462). A more recent study showed that hydrocortisone treatment can increase the ratio of long to short for PRLR expression in the mammary gland of ewes (463).

Recent studies indicate that thyroid hormone mediates isoform-specific expression PRLR in the mammary gland (464). Moreover, PRL isoforms are differentially regulated in the rat prostate by estrogen and testosterone (465).

Section VI. Biological Functions of PRL

PRL has been reported to regulate well over 300 functions in numerous target tissues, which can be generally categorized as reproductive, osmoregulatory, immunoregulatory, and metabolic. The major functions regulated by PRL, with an emphasis on those that pertain to this thesis will be described in this chapter.

A. Mammary gland development

The mammary gland is the primary target tissue of PRL, where it mediates mammary gland development (mammogenesis), synthesis of milk (lactogenesis), and maintenance of milk secretion (galactopoiesis). The role of PRL in mammogenesis and galactopoiesis will be

70 discussed in this section, while its function in regulating lactogenesis will be described later in this chapter.

The rodent (mouse) mammary gland develops in four stages, 1) in utero, (prenatal) where a ductal structure is produced, 2) during puberty, when the ducts elongate and branch to fill the mammary fat pad, 3) during the estrous cycle, when ductal side branches and alveolar buds develop, and 4) during pregnancy, when the ductal branches further elongate, and the alveioli differentiate into lobuloalveolar buds, becoming capable of producing copious amounts of milk. In most species, PRL, along with estrogen, progesterone, insulin, glucocorticoids, and GH are the principle hormones required for the development and differentiation of the mammary gland (401). While the combination of factors involved varies among species, PRL is a common requirement. It plays a vital role during lactation in rodents, rabbits and humans, and is less important in some ruminants (466).

The role of PRL in each developmental stage has been investigated thoroughly using in vitro and in vivo studies, and more recently through a detailed analysis of PRL -/- and PRLR -/- mice. Using cultured mammary epithelial cells, Ceriani et al. determined that embryonic development of the mammary gland does not require the presence of ovarian or pituitary factors (467). Furthermore, at birth, both PRL -/- and PRLR -/- mice possess a mammary ductal structure identical to wild-type mice (468;469). The changing hormonal environment at the onset of puberty is the controlling factor for the sexually dimorphic growth and development of the mammary gland. To evaluate the factors regulating these processes, early studies used hormonal replacement therapy in hypophysectomized, ovariectomized, and adrenalectomized mice (470). In these studies, the development of the mammary ducts was induced by a combination of estrogen and GH, while in males testosterone inhibited further growth of the mammary gland (471). Ormandy and Horseman made similar observations in both PRLR -/- and PRL -/- females, but noted that ductal side branching was reduced compared to wild-type mice (468;469). Similar effects were observed in Stat5a -/- mice, indicating that PRL mediates its effects in the mammary gland primarily via the Jak2/Stat signaling pathway (435).

By adulthood the complete ductal architecture of the gland has been established. Fully differentiated alveolar buds exist at the ends of major ducts and side branches of wild-type females, but not in PRL -/- or PRLR -/- mice. This demonstrates the importance of PRL in directing the differentiation of alveolar buds in the developing mammary gland. Interestingly, PRL exerts these effects via indirect mechanisms, as mammary PRLR, whether in the epithelium or the stroma, is not required for normal ductal side branching to occur (472).

During early pregnancy, elevated levels of serum PRL induce lobuloalveolar development in preparation for lactation. As PL levels rise around mid-gestation, mammary gland development becomes independent of pituitary PRL (473). PRL -/- and PRLR -/- mice are infertile, preventing analysis of the effects of pregnancy on mammary gland development. PRLR +/- mice, which are fertile, fail to lactate after the birth of their first litter due to stalled mammary gland development during late gestation, but regain their capacity lactate following subsequent pregnancies (468).

Involution of the mammary gland, which is induced by weaning, is a two step process, characterized first by engorgement, then by programmed alveolar cell death. Suckling can

71 prevent the onset of the second phase of involution, indicating PRL may be involved (474). Furthermore, serum PRL levels fall after weaning, as does Stat5 phosphorylation in the mammary gland (475). Capuco et al. reported that pregnancy inhibits mammary cell apoptosis after weaning while permitting proliferation of the mammary epithelium. Moreover, the authors observe that pregnancy also blocked the loss of Stat5a phosphorylation during involution, leading them to hypothesize that Stat5a and progesterone-signaling pathways act in concert to mediate this effect (476). Finally, Travers et al. have demonstrated that administration of PRL can prevent the cell death associated with involution (477).

B. Luteal Function

The corpus luteum, which represents a transformed follicle after ovulation, is an endocrine organ that synthesizes and secretes progesterone. Progesterone is necessary for implantation of the fertilized ovum, the maintenance of pregnancy, and in inhibition of ovulation. The PRLR is present in this tissue, and PRL is a well characterized regulator of luteal function, mediating actions that vary across species and depend on reproductive status (478). Early studies in rodents indicated that PRL acts as a luteotrophic factor by maintaining the structural and functional integrity of the corpus luteum. Indeed, it is now known that PRL is required for normal function of the corpus luteum throughout pregnancy in rodents. Following ovulation on the morning of estrous, the corpus luteum produces progesterone for two days, and then involutes if mating does not occur. In case of mating, the corpus luteum is rescued by twice daily surges of PRL for 10-12 days, as described earlier in this chapter. After pituitary PRL surges cease, the corpus luteum is maintained for the duration of the pregnancy by PL (479).

The luteotrophic actions of PRL are characterized by enhanced progesterone production, which is induced by permitting other luteotrophic hormones to act, as well as by acting directly to stimulate steroidogenesis. For instance, in the absence of PRL, progesterone is converted by 20αhydroxysteroid dehydrogenase to an inactive metabolite, 20αhydroxyprogesterone. PRL inhibits the activity of this enzyme, thereby increasing levels of active progesterone (480). PRL is also affects numerous genes whose expression is vital to the function of the corpus luteum. Notably, PRL increases expression of luteinizing hormone receptor (LH-R), and subsequently LH binding, which acutely stimulates progesterone production (481;482). LH, however, is not required for the maintenance of the corpus luteum. Gibori et al. suggest that PRL and estradiol synergize to elicit maximal progesterone secretion, possibly via PRLR/Jak2/Stat5 mediated up- regulation of estrogen receptor (ER) α/β. While the long PRLR isoform is expressed at higher levels in the corpus luteum, these authors also reported that the intracellular domain of the short form binds a luteal isoform of 17beta hydroxysteroid dehydrogenase 7, named PRLR associated protein, or PRAP (483).

C. Osmoregulation

PRL is viewed as an essential regulator of water and electrolyte balance in freshwater and euryhaline fish exposed to a freshwater environment. To survive in this setting, fish must prevent the loss of ions to the external hypoosmotic surroundings, and prevent the influx of water (484). PRL reduces ion loss and water permeability of osmoregulatory surfaces including the gill, kidney, intestine, urinary bladder (485-487). Fish PRLR are structurally similar to those of

72 higher vertebrates, but differences exist between teleosts and nonteleosts in their tertiary structure. The teleost PRLR, which most resembles the mammalian long form, varies in its intracellular domain between species, and is expressed primarily in the gill, kidney and intestine (488). Limited studies in one fish species have demonstrated activation of the Jak2/Stat5 pathway, but it is still not known if other signaling cascades also activated (489). There is little information on the cellular and biochemical effectors of the osmoregulatory actions of PRL (490). Herndon et al. reported that PRL inhibited the development of chloride cells, which are necessary for survival in a saltwater environment, while Pisam et al. observed an increase in the number of ion uptake cells in PRL treated tilapia (491;492). In many fish, freshwater exposure increases PRL expression, synthesis, secretion (484). Plasma factors such as osmolality and cortisol levels appear to mediate PRL secretion in tilapia (490). PRL also mediates osmoregulatory processes in birds where it eliminates excess salt through the nasal salt gland, a process which is stimulated by PRL (493). In mammals, PRL has been shown to reduce renal Na+ and K+ excretion and to stimulate Na+-K+ ATPase (494). Furthermore, PRL decreases Na+ and Cl- in sweat and increases water and salt absorption in the intestines (495;496).

D. Immunoregulation

PRL, which is structurally related to immunoregulatory factors, such as erythropoietin, granulocyte-macrophage colony stimulating factor and the interleukins, has long been implicated in immune function. Baroni et al. noted atrophy of the thymus and lymphoid organs, as well as abnormal immune responses in Snell dwarf mice, which lack PRL, GH, and thyrotropin (497). Subsequent studies determined that both PRL and GH were modulators of antibody production, mitogen-induced lymphocyte proliferation, as well as splenocyte and thymocyte number and cellularity (498). Bromocriptine treatment in rats led to attenuation of humoral or cell-mediated immunity, which was reversed by PRL treatment (499). Berczi et al. noted that PRL or GH administration into these rats caused an increase in the weight of the spleen and thymus (500).

One way by which PRL modulates immune system function is by stimulating mitogenesis and cell survival. Viselli et al. reported that normal T lymphocytes proliferate in response to PRL treatment (501). Furthermore, antibodies to PRL inhibit mitogen-stimulated lymphocyte proliferation in vitro (502). PRL also induces mitogenesis in Nb2 cells, a rat cell line derived from T lymphocytes (503). Given their dependence on the mitogenic activity of PRL, Nb2 cells are used as a very sensitive bioassay for PRL. Conversely, some studies have reported that PRL decreases mitogen stimulation of thymocytes. Murphy et al. observed decreased thymic cellularity in PRL treated dwarf mice, while no changes were observed in normal mice (504). Furthermore, treatment with PRL antiserum increases the number of cells in the thymus and spleen of young mice (505). It has been suggested that the thymopoietic effects of GH predominate over those of PRL, and that the effects of PRL on T cells may depend on their stage of differentiation (506).

PRL is globally seen as an immunostimulatory factor. Administration of PRL is associated with graft rejection in patients receiving heart transplants (507). In addition, PRL activates macrophages and increases superoxide anion production (508). Woody et al. propose the use of PRL to stimulate T cell production and function in immunocompromised patients, to augment thymopoiesis in aged individuals, and to accelerate T cell recovery following bone marrow

73 transplantation (509). PRL often acts in concert with other factors, serving as a coactivator of immune function. For example, Clevenger et al. report that PRL acts as a progression factor during IL-2 stimulated lymphocyte proliferation (510). Recent observations in patients with burn injuries revealed a positive correlation between serum PRL and levels of several interleukins, including IL-6, IL-8, and IL-10 (511).

The question of whether PRL is involved in the development of the immune system was recently examined using PRL -/- mice. Horseman et al. reported normal hematopoiesis, with no deficiency in the number of B lineage cells in the bone marrow, or CD4- and CD8-expressing cells in the thymus (469). Furthermore, they observed a normal distribution of B and T cells in the secondary lymphoid organs of PRL -/- mice. These results suggest that PRL is not critical for development hematopoietic system. Krishman et al. noted reduced glucocorticoid-induced thymocyte apoptosis in wild-type mice as compared to PRL -/- mice, suggesting that elevated PRL functions to maintain survival and function of T-lymphocytes (512).

E. Behavior

PRL is involved in the regulation of parental behavior of numerous species, including fish, birds, and mammals. Early evidence came from observations in fish, where PRL was implicated in fin fanning, an action that provides a constant supply of water to the eggs (513). Ogawa et al. reported that PRL stimulates the production of mucus used to feed young goldfish (514). In some species, PRL induces migration. Smith et al. reported that the migration from saltwater to freshwater of some teleosts was mediated by PRL (515). More recent studies agree with this, reporting that the PRL gene is elevated during spawning migration of salmon (516). Furthermore, injection of PRL into salamanders induced their migration from land to water (517).

In birds, PRL maintains readiness to incubate by inducing a quiet sitting behavior (518). PRL administration in laying turkey hens causes a sudden onset in incubation behavior, defined as an increase in nest visits, which can be reversed by PRL immunoneutralization (519;520). In addition, these hens also displayed a gradual decrease in egg laying during the time they were receiving PRL treatment, another indicator of the onset of incubation. March et al. also reported that immunization of chickens against PRL reduced the incidence or delayed the development of incubation behavior (521). In three types of albatrosses, high PRL levels were characteristic of the entire egg incubation period with a significant decline in concentrations towards the end of the brood-guard period (522).

In mammals, PRL has long been implicated in maternal behavior. Generally, high levels of PRL present at the end of pregnancy and throughout lactation mediate these actions. Voci et al. first observed enhanced pup retrieval and nest building following administration of PRL and progesterone near the preoptic region of the hypothalamus in mice (523). Bridges et al. subsequently reported that administration of PRL, or treatment with ectopic pituitary grafts stimulated maternal care in inexperienced, hypophysectomized female rats (524). Ovariectomy eliminated PRL-induced maternal behavior in the aforementioned animals, indicating the importance of steroids in regulating these actions (525). These, and later experiments identified both the mPOA and lateral hypothalamus as the hypothalamic regions where PRL mediates

74 maternal behavior (526). Interestingly, Anderson et al. recently reported that reproductive experience increases central PRL responsiveness via long-term up-regulation of PRLR in the medial preoptic area (527).

Observations in PRLR-deficient mice highlight the importance of PRL in regulating maternal behavior, as both PRLR -/- and PRLR +/- nulliparous females show a deficiency in pup retrieval and crouching behavior. Moreover, primiparous heterozygous females exhibit a profound deficit in maternal care when challenged with foster pups (528). Lucas et al. have proposed that PRLR +/- deficiencies occur because a threshold level of PRLR signaling is necessary for the induction of maternal behavior.

F. Prostate

The prostate is a key gland in the sexual physiology of male mammals. Its location in the reproductive tract influences several vital functions related to micturition, seminal emission and ejaculation. Androgens are the most important regulators of the prostate, but there is considerable evidence that PRL plays a role in prostate growth and function. In 1982, Prins et al. reported that increased serum PRL levels produced by pituitary grafts specifically enhanced testosterone-stimulated growth of the rat lateral prostate (529). Furthermore, the authors observed a biphasic effect of PRL on prostate growth, with stimulatory effects at lower doses, and no effects at high levels (530). These authors later demonstrated that PRL indirectly regulates lateral prostatic growth by increasing nuclear androgen receptor levels, thus optimizing its response to testosterone (531).

PRL can also directly regulate prostate growth. In vitro studies have shown that PRL is mitogenic in rat dorsal and lateral prostate (532). Ahonen et al. report that PRL is a survival factor for androgen-deprived rat dorsal and lateral prostate epithelium in organ culture (533). Furthermore, expression of both PRL and the PRLR in the prostate is increased by androgen treatment in vivo, and PRLR level is also increased by PRL itself (534;535).

Several recent in vivo observations support PRL as a regulator of prostate growth. For instance, transgenic overexpression of PRL resulted in stromal hyperplasia and intraepithelial dysplastic features within the prostate (536). In PRLR -/- mice, there is a small increase in dorsolateral and ventral prostate weight but no change in the weight of the anterior prostate (537). The dorsal but not ventral or lateral lobes showed a 12% loss of epithelial cells, with no other apparent changes in prostate morphology. There are likely compensatory mechanisms in place that account for the lack of phenotype in PRLR-deficient mice. Interestingly, Stat5a deficiency was associated with a distinct prostate morphology, characterized by increased prevalence of local disorganization within acinar epithelium of ventral prostates (538).

Section VI. Involvement of PRL in metabolic homeostasis

A. Body weight, adiposity, and food intake

The effects of PRL on body weight, adiposity, and food intake have been studied in a number of species. Studies in doves first demonstrated the orexigenic and weight stimulating affects of

75 PRL. While investigating the effects of PRL on incubation behavior, Buntin et al. observed that subcutaneous or peripheral injections of PRL markedly increased food intake and body weight in both male and female ring doves (539-541). Later studies determined that central administration directly into the VMN induced the most robust feeding response, with lesser effects observed when PRL was injected into the mPOA (542). Since hyperphagia induced by microinjections of PRL into the VMN was attenuated by coinjection of PRLR antibodies, it was speculated that is a primary site of PRL action in promoting hyperphagia in this species. However, other diencephalic loci, such as the mPOA and tuberal hypothalamus (TU), may also contribute to the orexigenic action of this hormone (543). Interestingly, later studies demonstrated that VMN destruction strongly perturbed feeding and body weight regulation in doves, but did not eliminate PRL-induced hyperphagia (544). This suggests that other diencephalic sites mediate the stimulatory effects of PRL on feeding when VMN function is impaired.

Indeed, more recent studies indicate a role for the orexigenic factors NPY and AgRP in PRL- induced hyperphagia. In addition to increasing food intake, i.c.v. administration of PRL significantly elevates AgRP-immunoreactive cell numbers in the TU, and NPY-immunoreactive cells in the infundibular region of non-breeding doves (545;546). It is hypothesized that rising PRL secretion during late incubation and early post-hatching initiates this increase in AgRP- and NPY-producing cells to regulate food intake and meet increasing energy demands.

The most consistent data linking PRL with body weight and food intake in mammals comes from studies in rats. In 1986, Moore et al. demonstrated that hyperprolactinemia, induced by ectopic pituitary explant under the kidney capsule, stimulated increased food intake and weight gain in female rats (547). Later studies confirmed these results, demonstrating that peripheral injection of PRL also increased food consumption and weight gain in a dose-dependent manner (548;549). Moreover, these experiments revealed that PRL exerted its effects in the absence of progesterone, which is known to increase food intake, weight gain, and adiposity (548;550).

The site at which PRL exerted its effects was unclear until Noel et al. reported that although peripheral administration of PRL increased food intake and body weight, i.c.v. administration increased food intake to the same extent without affecting weight gain (551). These data suggested that PRL acts both peripherally and centrally to regulate energy balance in female rats. Sauve et al. verified these observations, reporting that these effects also occur independent of ovarian steroids (552). It has since been shown that PRL significantly increases food intake when injected into the PVN, while VMN injections result in a non-significant trend towards a hyperphagic response (553). While these experiments have revealed at least one brain site at which PRL acts to increase food intake, they do not rule out the possibility that the effects of PRL on food intake may also involve other brain areas.

Surprisingly, the sex-specific effects of PRL gained little attention until 1999, when Heil et al. reported that food intake was unchanged in male rats treated with PRL (554). The author later reported that genetic male rats organized as females by postnatal castration significantly increased their food intake, while genetic female rats organized as males by postnatal androgen treatment maintained baseline levels of food intake. Their data indicate that organizational, but not activational, gonadal hormone exposure plays a critical role in the development of this sex- specific response to PRL (555).

76

There is a minor controversy as to whether PRL actually increases food intake and body weight in female rats. In 1991, Adler et al. reported that weight gain and food intake were the same in AP-grafted rats and in control muscle-grafted rats. Furthermore, they found that injection of PRL did not increase weight gain in normal or hypophysectomized rats. Adler concludes that neither chronic PRL excess caused by AP grafts nor acute PRL excess caused by PRL injections increases food intake or weight gain in female rats (556). Despite this report, the majority of available information is in support of PRL as a positive regulator of food intake and weight gain in female rats.

In humans, sustained elevation of PRL is occasionally accompanied by increased weight. Though it was not the focus of an early report, PRL levels and body weight were positively correlated in women by Wang et al. in 1987 (557). Other studies noted that some women with a history of recent weight gain also had higher PRL levels, albeit still in the normal range, as compared to control women without such a history (558). Greenman et al. and several others have reported increased body weight associated with PRL secreting pituitary adenomas, an effect that was reversed by bromocriptine treatment in 90% of subjects. Interestingly, men were significantly more responsive than women to this treatment (559-561).

These observations led to the hypothesis that elevated PRL levels are a potentially reversible cause of weight gain. However, this is highly debated for several reasons. First, bromocriptine induced weight loss is usually modest and delayed, and second, obesity is present in less than 30% of prolactinoma patients, and that only 25% of these lose weight when treated with bromocriptine (562;563).

While there is a correlation between increased PRL levels in patients taking neuroleptics (D2R antagonists) and weight gain, recent studies in rats indicate that weight gain may occur independent of changes in PRL levels. In a putative animal model of antipsychotic drug-induced weight gain, female rats treated with low doses of olanzapine or resperidone increased food intake and gained weight without elevating serum PRL levels (564;565). Furthermore, in male rats, sulpiride administration did not affect bodyweight gain and food intake despite significantly increasing PRL (566). In humans, it remains unclear whether PRL is responsible for weight gain observed in patients treated with antipsychotics, as different drugs appear to exert different effects. Clozapine reportedly increases weight gain in humans while having little effect on serum PRL, whereas a significant positive correlation was observed between PRL levels and body weight gain in subjects treated with sulpiride (567). Baptista et al. reported that PRL and BMI correlated positively in men but not women treated with antipsychotics (568;569). Because many of the atypical antipsychotics have a broad spectrum of action as neurotransmitter receptor antagonists, affecting serotonin, dopamine, adrenoceptors, and histamine receptors, it is difficult at this point to separate the effects of PRL from other alterations in brain circuitry (570). There is presently no strong evidence to indicate that, at normal circulating levels, PRL is a major factor in human obesity (571).

Limited and controversial information regarding PRL and food intake/weight gain has been reported in mice. Early studies reported that suppression of PRL by bromocriptine treatment increased food intake, but had no affect on weight gain in females (572). There were no

77 experiments performed in males in this study, so the sex-specific effects of PRL suppression remained unknown. Later, in a model of hyperprolactinemia, male mice receiving ectopic pituitary grafts exhibited small, but significant increases in food intake and body weight (573). Furthermore, serum FFAs and the weight of epididymal fat bodies were significantly lower in hyperprolactinemic mice. In female transgenic PRL-overexpressors, a minor increase in body weight was observed, while no differences were found in adipose tissue (450). Whereas these data tend to suggest that PRL is a positive stimulator of food intake and body weight, the effects are minor at best, and are derived from very limited observations. Several recent studies have attempted to identify the role of PRL in weight gain and adiposity using PRLR-deficient mice. In 2001, Freemark et al. reported that female PRLR-/- mice grew at a normal rate until 16 weeks of age, at which time a small (5-12%), but progressive reduction in body weight developed in both sexes. These authors reported that females were affected to a greater extent than males, and that the reduction in weight gain was associated with a marked 50% reduction in total abdominal fat mass (4). In a study published one year later, the same authors reported that 16-week old PRLR- /- females weigh 25% (5 grams) less than their wild-type littermates, while male PRLR-/- mice grew at a normal rate. Finally, in 2005 Fleenor et al. reported no differences in body weight of either male or female PRLR-/- mice when compared to wild-type littermates (574). Interestingly, although it was not the focus of an early study of PRLR-/- mice, Lucas et al. noted in 1998 that body weights of PRLR -/- and wild-type mice showed no differences (528). Unfortunately, in each of these studies, the authors failed to cite their own previous publications. Moreover, they did not mention that these mice develop macrodenomas with age, likely resulting in altered pituitary function (575).

B. Lactogenesis

During lactation, nutrients are redirected from adipose tissue stores to the mammary gland, so as to provide large amounts of energy as milk for the nursing young. The epithelial cells of the alveoli, which synthesize and secrete milk ultimately determine the quantity and quality of milk production.

It is difficult to assess the direct actions of PRL on mammary gland metabolism since there are no fully functional in vitro models of the lactating mammary gland. Therefore, most of our knowledge on the effects of PRL on lactation is inferred from PRL suppression in vivo either by bromocriptine treatment or litter removal. Hence, in vivo models can be problematic because it is difficult to distinguish direct vs. indirect actions of PRL.

All three major constituents of milk, e.g., proteins, lactose, and lipids, are regulated by PRL. The milk protein component is broadly subdivided into two fractions, casein and whey proteins (576). Casein is the predominant phosphoprotein found in milk, and due to its hydrophobic nature, it exists as a suspension of particles called casein micelles that are held together by Ca2+ ions and hydrophobic interactions. PRL has long been reported to induce transcription of β- casein expression via Stat5 binding to its promoter and enhancer (577). Optimal β-casein expression is achieved via stimulation by both PRL and glucocorticoids, which according to a recent study, requires synergistic formation of a multi-protein complex at β-casein regulatory regions (578).

78 Whey is the remaining liquid in milk after it has been curdled and strained. It is rich in globular proteins known as whey proteins. These include whey acidic protein (WAP), as well as α- lactalbumin and β-lactobumin. WAP is differentially induced across species by the synergistic actions of several factors. For example, in mammary gland cultures from rat, mouse, and pig, WAP gene expression requires insulin, cortisol and PRL. However, in rabbits, WAP gene expression is induced by PRL alone, but increases further when insulin and cortisol are added (579). Mutational studies have identified a recognition site for Stat5a, which mediates PRL induction of WAP expression (580).

Lactose is the major carbohydrate in milk, being composed of galactose and glucose. α- lactalbumin, which constitutes only about 2% of the whey fraction of milk, is also a component of the lactose synthase complex that uses glucose and UDP-galactose as substrates for the synthesis of lactose in the Golgi of mammary epithelial cells. Oppat et al. have demonstrated that PRL increases the synthesis of lactose in cultured mammary glands by inducing glucose uptake and α-lactalbumin availability (581).

Lactation results in a major re-adjustment of lipid metabolism to meet the demands of the mammary gland for milk fat synthesis. This includes a large increase in mammary gland TG synthesis from circulating fatty acids or via de novo lipogenesis, and a concomitant decrease of these processes in adipose tissue (582). TG comprise up to 98% of the lipid content of milk, reaching up to 50% of the total milk volume. Changes in the fatty acid composition of the diet are directly reflected in milk composition. The effects of total food withdrawal on milk lipid synthesis depend on the species and the duration of fasting. In rodents who have little metabolic reserve in comparison to the rate of milk synthesis, 24 hr of food deprivation leads to a state of starvation, with profound effects on the rate of synthesis of all milk components, including lipids (583). On the other hand, hibernating species fast during lactation without affecting milk secretion (584).

PRL enhances lipid production in the mammary gland by increasing the expression and activity of lipogenic enzymes, such as ACC and FAS (585-587). Furthermore, PRL prevents phosphorylation-mediated inactivation of pyruvate dehydrogenase, thereby increasing acetyl- CoA availability (588). Collectively, these alterations in enzyme activity and expression enhance the storage of lipids in the mammary gland to meet energy demands during lactation. Notably, although PRL levels decrease as lactation is established, nursing stimulates pituitary PRL release thereby promoting continued milk production.

PRL was originally identified as the major regulator of mammary LPL, since the activity of this enzyme was significantly decreased 24 hr after hypophysectomy or by sealing of the teats, with some effects reversed by PRL injections (589). However, more recent experiments demonstrated that bromocriptine alone had little effect on rat mammary LPL activity and expression, although bromocriptine combined with anti-GH serum produced a marked reduction in both (590). This finding is consistent with observations that PRL deficiency did not alter triolein uptake into the lactating mammary gland (591). Furthermore, in this study, injection of PRL did not overcome the effects of insulin and PRL deficiency on triolein uptake or LPL activity in the lactating rat mammary gland, whereas insulin treatment did. These authors conclude that insulin is the primary regulator of TG uptake as well as LPL activity in the lactating mammary gland of the rat

79 and that the action of PRL on these processes is indirect (591). A study by Vernon et al. agrees, reporting that PRL influences the rate of fatty acid synthesis by modulating the insulin-binding capacities of the tissues (582). Insulin tends to increase mammary LPL activity in rats, indicating that it stimulates lipogenesis (592). Da Costa et al. showed that insulin deficiency decreased LPL activity and the uptake of labeled triolein by rat mammary gland in vivo, an effect that was partially reversed by exogenous insulin (591).

C. Adipokine release

Prior to the detection of PRLR in adipose tissue, PRL was considered an indirect regulator of adipose tissue function. For instance, early reports suggested that lactogens exert lipolytic effects in WAT during pregnancy and lactation since elevated PRL during lactation is accompanied by a reduction of abdominal fat stores (593). The recent discovery of PRLR in adipose tissue indicates direct actions of PRL and has generated a great deal of interest in the effects of PRL on this tissue. Currently, in vivo studies addressing the effects of PRL on adipose tissue function have employed mouse models that either over-express or lack PRL. Adipose tissue explants, primary preadipocytes, mature adipocytes, and adipocyte cell lines are also used to study the effects of PRL in vitro.

Circulating levels of adiponectin decrease significantly throughout gestation in both female mice and in humans, suggesting that PRL suppresses adiponectin secretion (594). Indeed, Combs et al. reported that PRL treatment in mice decreased adiponectin levels, while bromocriptine had the opposite effect (595). Interestingly, in transgenic mice overexpressing adiponectin, PRL levels are significantly (2-fold) elevated, while GH, leptin, and glucocorticoids are similar to those in wild-type mice (191). These authors suggest that the elevated PRL in adiponectin overexpressing mice serves as a compensatory mechanism for down-regulating adiponectin. Notably, PRLR deficiency has no effect on adiponectin levels in either males or females, leading to speculation that the suppression of adiponectin is a result of direct hormonal regulation (182). In the same study, adiponectin levels were decreased by ~50% in PRL overexpressing females, but were unchanged in males with elevated PRL. It is possible that the modified adiponectin levels are a result of sex-specific hormonal regulation since in wild-type mice, adiponectin levels in males are approximately one-third of the levels found in females.

The relationship between PRL and adiponectin is not entirely clear however. Serum adiponectin is negatively correlated with adiposity, but PRL-overexpressing mice have decreased fat and lower adiponectin levels than wild-type mice (182;450). Nilsson et al. believe that reduced adiponectin in PRL-overexpressing mice are reflective of insulin sensitivity, rather than adiposity. Perhaps so, as PRLR deficient mice show no differences in adiponectin and have normal peripheral insulin sensitivity, but insulin sensitivity has not been measured in the PRL- overexpressors.

Limited in vitro studies support the notion that PRL inhibits adiponectin secretion. In human subcutaneous abdominal adipose tissue explants, PRL decreased adiponectin secretion, while increasing the expression of AdipoR1, but not AdipoR2 (182). A recent study by Asai-Soto confirmed these results using isolated primary adipocytes, while the changes in receptor expression have not been confirmed (594). It has been proposed that there is an

80 autocrine/paracrine loop for adiponectin regulation in adipose tissue, but it is unclear why adiponectin secretion is down-regulated while its receptor is up-regulated.

Although leptin is the most-studied adipokine, a clear connection with PRL has not been established. Serum leptin levels are markedly elevated in PRL-overexpressing, pituitary grafted, or PRL treated mice (596;597). Conversely, PRLR-deficiency results in decreased leptin levels (4). While these observations suggest that PRL is a positive regulator of leptin secretion and/or expression, there is currently no explanation why elevated PRL leads to increased leptin levels. Increased leptin is not a reflection of changes in adiposity in these studies, since hyperprolactinemic mice do not have increased body fat. In fact, PRL-overexpressing mice have minor decreases in body fat (450). The decreased leptin observed in PRLR deficient mice was believed to be a result of reduced fat mass, with more recent studies reporting no differences in circulating leptin or fat mass (4;574).

Few in vitro studies have investigated the regulation of leptin by PRL. In cultured female mouse adipocytes, PRL alone had no effect on leptin, but inhibited leptin production in the presence of insulin (596). Conversely, in differentiated brown adipocytes, PRL potentiates the stimulatory effect of insulin on leptin release (598). The effects of PRL on leptin secretion are depot specific, and may require the presence of cofactors.

Finally, a recent study in our laboratory showed that addition of exogenous PRL to differentiated LS14 cells, a novel human adipocyte cell line, inhibits the expression and release of IL-6, a pleiotropic cytokine involved in inflammation and tissue remodeling (354).

D. Lipid Metabolism

Only limited data are available on the actions of PRL on lipid metabolism in adipose tissue (Fig. 8). Early observations suggested that the hyperprolactinemia of lactation was responsible for the blunting of both storage and mobilization of lipids from abdominal adipose tissue to the mammary gland (599;600). For instance, Zinder et al. noted that when lactating rats were hypophysectomized, LPL activity increased in adipose tissue and decreased in mammary gland, an effect that was reversed by PRL treatment (589). Furthermore, Agius et al. found that removal of the suckling stimulus or treatment with bromocriptine significantly increased lipid storage in adipose tissue. Again, this effect was reversed by PRL injection (601). Taken together, the available data indicated that PRL inhibits lipogenesis, increases lipolysis, or performs both actions in adipose tissue. However, there is one report questioning whether PRL is the primary factor regulating the mobilization of lipids during lactation. A study by Oller do Nascimento et al. reported that bromocriptine treatment did not alter lipid accumulation or lipogenesis in adipose tissue (593). Moreover, the authors stated that PRL treatment did not alter the response seen in litter-removed lactating rats. While the explanation for these results remains unknown, the authors’ reasoning that the lack of PRLR in adipose tissue was a cause is no longer valid. On this note, numerous studies have been conducted over the years in an attempt to elucidate the actions of PRL on lipid mobilization in adipose tissue. While the recent discovery of the PRLR in adipose tissue has reinvigorated interest in this topic, the actions of PRL remain controversial and relatively unknown.

81 PRL Leptin Glucose P Jak2 Jak2 P Adiponectin

Y P Stat5 P P IRS Stat5 Resistin Glut4 ? IL-6 P Stat5 Stat5 P P ? FIAF LPL

FAS Malonyl CoA VLDL ? FFA HSL FFA FFA Triglycerides

G3P Glycerol Glycerol

Figure 8: A model summarizing known and putative actions of PRL on adipocytes. The activated PRLR signals primarily through the Jak2/Stat pathway, but is also known to activate IRS and Akt. PRL prevents lipid influx/lipogenesis by inhibiting lipoprotein lipase (LPL) and fatty acid synthase (FAS). In addition, PRL reduces the secretion of adiponectin and IL-6, either stimulates or inhibits leptin, and has no effect on resistin. The effects of PRL on hormone sensitive lipase (HSL), glucose transporter 4 (GLUT4), and fasting-induced adipocyte factor (FIAF) secretion are currently unknown.

82 Only few studies focused on potential direct effects of PRL on lipogenesis in adipose tissue. For instance, Flint et al. determined that reduction of PRL during lactation via bromocriptine treatment or pup removal led to increased activity of the lipogenic enzymes FAS and LPL in adipose tissue (587). This agrees with the inhibition of LPL activity in human adipose tissue explants (353). Moreover, PRL at a very high dose down-regulates FAS expression in 3T3-L1 adipocytes via a Stat5a mechanism (602).

Despite numerous reports, it also remains unclear whether PRL is lipolytic. Most of the available data indicate that PRL does not exert lipolytic activity. For example, early studies reported that PRL had no effect on lipolysis in fat explants from rabbits and non-lactating ruminants (603;604). On the other hand, Williams et al. reported that PL was lipolytic in adipose tissue from both pregnant and non-pregnant women (605). Other studies have shown that PL stimulates lipolysis in adipocytes isolated from rats (606;607). A criticism of these early studies is that the PRL or PL used may have been contaminated with trace amounts of other hormones, such as GH (608). In later studies, Fielder et al. reported that PL had no effect on lipolysis in adipose tissue from virgin or pregnant mice, while PRL only induced glycerol release at a superphysiological concentration (5μg/ml) (609). Notably, others reported that PRL was devoid of lipolytic activity in several species, including rats, mice, hamsters, guinea pigs and rabbits (603). Taken together, most available data indicate that PRL is not lipolytic. Given that only one study using human tissue was reported, more experiments should be performed to determine whether this phenomenon is real or not.

Interestingly, Carbrera et al. observed that adipocytes isolated from hyperprolactinemic rats had decreased sensitivity to the anti-lipolytic/lipogenic effect of insulin, despite increasing levels of insulin receptors (610). This observation is similar to that seen in adipocytes from lactating rats, which show poor responsiveness to insulin in terms of lipid synthesis from glucose (611). In contrast, insulin significantly increases lipid synthesis in adipocytes from bromocriptine treated rats, while PRL administration partially reversed the effects of bromocriptine (610). These observations led Cabrera et al. to speculate that PRL mediates adipocyte resistance to insulin during lactation. In agreeance with this, another study reported that PRL treatment decreased glucose transport in rat adipocytes, without altering insulin binding (612). On the other hand, PRL decreases insulin binding to adipocytes from women at term gestation (613). Taken together, these data indicate that PRL may participate in the tissue-specific insulin-resistance that occurs during pregnancy. However, a mechanism for this effect has not been elucidated.

PRL may also play a role in adipogenesis. PRLR expression is upregulated during differentiation of BMS2 cells, a bone marrow stromal cell line, 3T3-L1 adipocytes and T37i brown adipocytes (598;614). Moreover, PRLR mRNA levels rose 90-fold following induction of adipocyte differentiation in 3T3-L1 cells, the levels of PRLR mRNA rose 90-fold (615). Interestingly, expression of both long and short isoforms of the PRLR increased during differentiation, but the long isoform predominated at all time points.

For complete differentiation to occur, most murine preadipocytes require the presence of fetal bovine serum (FBS), which contains lactogenic hormones. Recent studies determined PRL and GH increase Stat5a and Stat5b activity in differentiating 3T3-L1 cells, and can replace the requirement for FBS during adipogenesis (615;616). Ectopic expression of Stat5a has also been

83 shown to confer adipogenesis in 3T3-L1 preadipocytes and in two different non-precursor cell lines (617;618). Additionally, Nanbu-Wakao et al. reported that PRL enhanced expression of C/EBPβ and PPARγ mRNA while stimulating the differentiaion of NIH-3T3 cells, probably via Stat5 mediated activation of the aP2 promoter (619).

In conclusion, the available data indicate that PRL may act as an adipogenic factor, while its effects on lipogenesis and lipolysis remain unclear. Furthermore, PRL appears to act in concert with insulin to exert some of its effects. Certainly, more research should be performed in this area, especially given the recent demonstration of PRLR signaling in adipocytes.

E. Pancreas

Lactogenic hormones are important regulators of pancreatic islet growth during pregnancy and the perinatal period. In the rodent pancreas, the PRLR is expressed primarily in acinar cells and ducts in early gestation. In late gestation and the postnatal period, the PRLR is expressed predominantly in pancreatic islets, co-localizing with insulin and glucagon. PRLR immunoreactivity increases significantly between embryonic days e16.5 and e18.5 in the fetal rat (620). These ontogenetic alterations in the distribution of PRLR expression support a role for lactogenic hormones in pancreatic growth and function during development. This notion is supported by studies in PRLR -/- mice, which as early as three weeks of age and through adulthood, exhibit reductions in islet size and density, as well as β-cell mass (621). In addition, the insulin content of islets is lower and the amount of insulin secreted in response to glucose injection is reduced when compared to wild-type mice. However, the insulin response is not impaired in these mice, since the clearance of glucose is normal when insulin is injected.

As mentioned earlier, pancreatic β cells undergo several adaptations during pregnancy to meet the increased demands for maternal insulin (247). Although the hormonal changes which occur during pregnancy are complex, lactogenic hormones appear to be sufficient to induce all of the up-regulatory changes that occur in islets during pregnancy in rodents. For instance, Sorenson et al. reported that infusion of PRL into rats decreased glucose stimulation threshold, enhanced insulin secretion, and increased beta cell coupling in islets (622). Notably, the changes induced by PRL closely resembled those observed in the islets of pregnant rats (623).

Several in vitro reports have confirmed and further characterized the importance of lactogens on β cell growth and function. In isolated islets from mice, rats, and chicks, PRL increases insulin secretion, cell-to-cell communication, and glucose sensitivity (624-626). These studies also demonstrated that PRL is more effective than GH in rodents but not in humans (247). Weinhaus et al. have shown that the increases in GK activity and GLUT2 levels that occur during pregnancy can also be induced by PRL treatment of islets in vitro (627). Similar PRL-mediated effects on GLUT2 were observed with INS-1 cells, although GK activity was not measured (628). Conversely, in a study using the glucose-responsive MIN6 β cell line, PRL increased GK activity without affecting GLUT2 protein levels (629).

The actions of PRL in the pancreatic ß-cell are mediated, at least in part by Stat5, which stimulates the expression of the genes for the PRLR, GK, and insulin (627;628;630). For example, continuous PRL stimulation induces a transient activation of Stat5a and a biphasic

84 activation of Stat5a in β cells, suggesting that prolonged elevation of lactogens during gestation support long-term activation of Stat5b in β cells (631;632). In contrast, they propose that pulsatile secretion of PRL during lactation is insufficient to maintain the up-regulation of islet function and may be part of the explanation of why they are at a lowered functional state during this period. PRL also induces nuclear migration of Stat5a and -5b in rat INS-1 cells and induces binding of INS-1 nuclear proteins to Stat5 sequences in the rat insulin-1 promoter (633). However, Fleenor et al. believe that the Stat5 motif may not be essential for PRL induction of insulin gene transcription because the magnitude of binding of Stat5 nuclear proteins was only 1/30th that of their binding to the ß-casein Stat5 site (634). Moreover, deletion of the Stat5 motif in the rat insulin-1 promoter has no effect on PRL induction of insulin gene transcription. Finally, PRL induces transcription of constructs containing the wild-type human and rat insulin-2 promoters, which have no classic Stat5-binding sequences. Notably, deletion of the rat insulin-1 promoter region containing the Stat5 binding sequences significantly reduces basal transcriptional activity (634). This finding suggests that this region is essential for full transcription of the insulin gene. Brelje et al. noted that PRL stimulated the nuclear translocation of Stat5b only in the insulin-containing β cells and not in the glucagon-containing α cells or the somatostatin-containing δ cells (632). These observations confirm that all islet cells have a functional Stat5 signaling pathway but that only ß-cells are PRL responsive.

PRL also regulates the structural and functional changes of the islets via pathways other than Jak2/Stat. Amaral et al. reported that in isolated rat islets, PRL induced a dose-dependent IRS-1 and IRS-2 phosphorylation, which was associated by increased PI3K activation (635). Furthermore, PRL also induced ERK1 and ERK2 phosphorylation in neonatal islets, demonstrating that PRL activates MAPK. Using a PRLR antisense oligonucleotide, this group later reported that downstream proteins of the PI3K (AKT and p70S6K) and MAPK (SHC and ERK1/2) cascades are regulated by PRL in islets from pregnant rats (441). Treatment with the PRLR antisense oligonucleotide also reduced glucose-induced insulin secretion in islets from pregnant rats. These findings indicate that PRL mediates the increase in islet mass and the sensitivity to glucose during pregnancy via the PI3K and MAPK pathways.

A recent micoarray analysis of PRL-treated rat islets revealed numerous altered genes (636). The differentially expressed transcripts were grouped into several categories including cell proliferation and differentiation, signal transduction, transcription factors and coactivators, translational machinery, Ca2+-mediated exocytosis, and immuno-response.

Interestingly, Sorenson et al. have shown that progesterone counteracts the effects of PRL on insulin secretion and β cell division in vitro (637). Moreover, the changes observed in islets under the influence of PRL and progesterone mimics those seen in islets during pregnancy. Another study reported that glucocorticoids inhibited PRL induced insulin secretion while also increasing β cell apoptosis (638). These data indicate that increasing levels of progesterone and glucocorticoids during late pregnancy could effectively reverse lactogen-induced changes in islet function by inhibiting insulin secretion and cell proliferation while increasing apoptosis.

In conclusion, it is clear that lactogens are important regulators of β cell growth and function, particularly during pregnancy in rodents. PL, present at high levels during pregnancy are likely the major lactogenic factors acting on the pancreas. On the other hand, the roles of both PRL and

85 PL in human pregnancy are unknown (248). The physiological relevance of PL secretion has been questioned because placentas of some species, e.g., pigs, dogs, and cats, do not produce a distinct PL (639). However, an increase in serum levels of PRL, may compensate for the absence of a distinct PL.

F. Liver

The pituitary exerts important regulatory effects on hepatic cholesterol and lipid metabolism in both animals and humans through the secretion of several hormones. Despite the expression of high levels of the PRLR, a major function for PRL has not been identified in the liver. PRL has been, however, linked to a number of enzymatic processes associated with bile synthesis, fatty acid oxidation and production, and LPL activity.

The secretory rate of taurocholate, a major conjugated bile acid, is increased in the livers of postpartum rats, suggesting PRL mediates this process. Indeed, this increase is blocked by bromocriptine, and is duplicated by PRL treatment (640;641). Studies by Liu and Ganguly have determined that PRL increases the synthesis and incorporation of the Na+- taurocholate transporter via Jak2/Stat5 (642;643). It is possible that the elevated production of bile during lactation aids in the digestion of the increased food ingested to meet energy needs.

Changes in the fatty acid profile are strongly correlated with hepatocyte membrane fluidity. Furthermore, biosynthesis of the fatty acids that are essential elements of biological membranes are largely modified by a wide variety of hormones, including PRL. Igal et al. demonstrated that hyperprolactinemia modifies the biosynthesis of PUFA and PUFA distribution in liver cell membranes, leading to increased lipid rotational mobility and fluidity (644). This observation agreed with an earlier report by Dave et al., of lower viscosity in hepatic membranes from PRL- treated rats (645). These authors also noted that a decrease in hepatic membrane viscosity is associated with a higher binding capacity for PRL, and that membranes with low fluidity possess a reduced number of PRL receptors.

There is some controversy as to whether PRL alters LPL activity in the liver. Machida et al. reported that LPL activity in the fetal liver treated with PRL was significantly higher than that of control. LPL activities in the rat fetal liver gradually increased until the time of birth, suggesting that PRL is one of the factors that regulate fetal lipid metabolism (646). Conversely, Julve et al. demonstrated that PRL decreases LPL activity in isolated hepatocytes from 5 day old rats (647). It is possible that the responsiveness of hepatocytes differs between prenatal and postnatal livers, but there are insufficient reports on this topic. One in vivo study reported that bromocriptine administration increased the rates of hepatic lipogenesis in females on day 21 of gestation, supporting the notion that PRL decreases LPL activity (648). Furthermore, removal of the suckling stimulus in lactating rats increases lipid production in the liver by 77%, an effect that is prevented by administration of PRL (601).

Other hepatic enzymes may be regulated by PRL. Most of those participate in energy generating or storing processes. For instance, Goodman et al. observed that PRL increased hepatic uptake of carnitine, which is responsible for the transport of fatty acids from the cytosol into the mitochondria where fatty acid oxidation occurs (649). In addition, suppression of PRL increases

86 the activity of cytosolic NADP-linked isocitrate dehydrogenase (ICDH), an enzyme which participates in the citric acid cycle. This effect was reversed by PRL injection. Short-term PRL incubation with isolated hepatocytes had no effect on the ICDH activity, suggesting that the effect of PRL is exerted at the transcriptional level (650). A study by de la Asuncion et al. has reported that hepatic uptake of amino acids was greater in bromocriptine-treated lactating rats and in lactating rats that had had their pups removed as compared with untreated lactating rats (651). These data are difficult to interpret. While it is assumed that the elevated PRL of lactation causes increased amino acid uptake, the elevation of amino acid uptake when PRL is removed makes it difficult to explain. It is possible that other factors present during lactation increase amino acid uptake, while PRL provides an inhibitory signal.

Taken together, these data indicate that PRL participates in various metabolic processes in the liver. Since there are only limited studies that have examined this issue, no definitive function has been ascribed for PRL in the liver. PRL likely acts in concert with other factors, and may play a part in the alterations of hepatic function that occur during lactation, including lipogenesis.

G. Prostate

The prostate gland of humans and other animals has the property of accumulating and secreting extraordinarily high levels of citrate. The prostate secretory epithelial cells synthesize citrate which due to a limiting mitochondrial aconitase, accumulates rather than being oxidized. Thus, citrate is essentially an end product of prostate metabolism. The key regulatory enzymes directly associated with citrate production in prostate cells are mitochondrial aspartate aminotransferase (mAAT), pyruvate dehydogenase, and mitochondrial aconitase. Testosterone and PRL are involved in the regulation of the genes associated with these enzymes. It is not known if the effects of PRL and testosterone are additive, synergistic, or independent. In addition, these hormones appear to exert differential effects in the distinct lobes of the prostate.

Franklin et al. have reported that PRL stimulates citrate production in the lateral prostate by increasing the transcription of mAAT (652;653). Recent studies by this group have determined that PRL mediates this effect via a PKC pathway (654). They have also have shown that PRL increases the expression of the E1 regulatory subunit of pyruvate dehydrogenase in rat ventral and lateral prostate cells; thereby regulating the availability of acetyl CoA for citrate synthesis (655).

Mitochondrial aconitase gene expression is increased by PRL in rat ventral prostate cells, pig prostate cells, and human malignant prostate cells (656;657). Liu et al. confirmed the observations made in rats, and further demonstrated that PRL increased citrate utilization and m- aconitase in ventral prostate cells, while having no effect in cells of the dorsal prostate (658). It has since been reported that in human prostate cells, this effect is mediated via a cAMP-mediated pathway (657).

The prostate gland of many animals also accumulates extremely high levels of zinc. There is good evidence that zinc is an important regulator of m-aconitase and citrate oxidation of prostate epithelial cells. It is known that prostate cells contain high levels of intracellular zinc, notably within mitochondria. Costello et al. report that the accumulation of zinc results in the inhibition

87 of mitochondrial aconitase activity which minimizes the ability of these cells to oxidize citrate (659). This is an important relationship associated with the unique functional and metabolic capability of the prostate to accumulate high citrate levels.

88 References

1. Mokdad AH, Ford ES, Bowman BA, Dietz WH, Vinicor F, Bales VS, Marks JS 2003 Prevalence of obesity, diabetes, and obesity-related health risk factors, 2001. JAMA 289:76-79 2. Eknoyan G 1999 Santorio Sanctorius (1561-1636) - founding father of metabolic balance studies. Am J Nephrol 19:226-233 3. Frankenfield DC, Muth ER, Rowe WA 1998 The Harris-Benedict studies of human basal metabolism: history and limitations. J Am Diet Assoc 98:439-445 4. Freemark M, Fleenor D, Driscoll P, Binart N, Kelly P 2001 Body weight and fat deposition in prolactin receptor-deficient mice. Endocrinology 142:532-537 5. Cinti S 2005 The adipose organ. Prostaglandins Leukot Essent Fatty Acids 73:9-15 6. NAPOLITANO L 1963 THE DIFFERENTIATION OF WHITE ADIPOSE CELLS. AN ELECTRON MICROSCOPE STUDY. Journal of Cell Biology 18:663-679 7. Geloen A, Collet AJ, Guay G, Bukowiecki LJ 1990 In vivo differentiation of brown adipocytes in adult mice: an electron microscopic study. Am J Anat 188:366-372 8. Christiansen EN, Pedersen JI, Grav HJ 1969 Uncoupling and recoupling of oxidative phosphorylation in brown adipose tissue mitochondria. Nature 222:857-860 9. Gregoire FM, Smas CM, Sul HS 1998 Understanding adipocyte differentiation. Physiol Rev 78:783-809 10. Casteilla L, Champigny O, Bouillaud F, Robelin J, Ricquier D 1989 Sequential changes in the expression of mitochondrial protein mRNA during the development of brown adipose tissue in bovine and ovine species. Sudden occurrence of uncoupling protein mRNA during embryogenesis and its disappearance after birth. Biochem J 257:665-671 11. Cannon B, Nedergaard J 2004 Brown adipose tissue: function and physiological significance. Physiol Rev 84:277-359 12. Clarke L, Bryant MJ, Lomax MA, Symonds ME 1997 Maternal manipulation of brown adipose tissue and liver development in the ovine fetus during late gestation. Br J Nutr 77:871-883 13. Lean ME 1989 Brown adipose tissue in humans. Proc Nutr Soc 48:243-256 14. Symonds ME, Stephenson T 1999 Maternal nutrition and endocrine programming of fetal adipose tissue development. Biochemal Society Transections 27:97-103 15. Poissonnet CM, Burdi AR, Bookstein FL 1983 Growth and development of human adipose tissue during early gestation. Early Hum Dev 8:1-11 16. Slavin BG 1979 Fine structural studies on white adipocyte differentiation. Anat Rec 195:63-72 17. Symonds ME, Mostyn A, Pearce S, Budge H, Stephenson T 2003 Endocrine and nutritional regulation of fetal adipose tissue development. Journal of Endocrinology 179:293-299 18. Youngstrom TG, Bartness TJ 1995 Catecholaminergic innervation of white adipose tissue in Siberian hamsters. Am J Physiol 268:R744-R751 19. Nnodim JO, Lever JD 1988 Neural and vascular provisions of rat interscapular brown adipose tissue. Am J Anat 182:283-293 20. Collins S, Surwit RS 2001 The beta-adrenergic receptors and the control of adipose tissue metabolism and thermogenesis. Recent Prog Horm Res 56:309-328

89 21. Hausman DB, DiGirolamo M, Bartness TJ, Hausman GJ, Martin RJ 2001 The biology of white adipocyte proliferation. Obes Rev 2:239-254 22. Shepherd PR, Gnudi L, Tozzo E, Yang H, Leach F, Kahn BB 1993 Adipose cell hyperplasia and enhanced glucose disposal in transgenic mice overexpressing GLUT4 selectively in adipose tissue. Journal of Biological Chemistry 268:22243-22246 23. Faust IM, Johnson PR, Stern JS, Hirsch J 1978 Diet-induced adipocyte number increase in adult rats: a new model of obesity. Am J Physiol 235:E279-E286 24. Sen A, Lea-Currie YR, Sujkowska D, Franklin DM, Wilkison WO, Halvorsen YD, Gimble JM 2001 Adipogenic potential of human adipose derived stromal cells from multiple donors is heterogeneous. J Cell Biochem 81:312-319 25. Gregoire FM 2001 Adipocyte differentiation: from fibroblast to endocrine cell. Exp Biol Med (Maywood ) 226:997-1002 26. Green H, Meuth M 1974 An established pre-adipose cell line and its differentiation in culture. Cell 3:127-133 27. Green H, Kehinde O 1979 Formation of normally differentiated subcutaneous fat pads by an established preadipose cell line. J Cell Physiol 101:169-171 28. Rosen ED, Walkey CJ, Puigserver P, Spiegelman BM 2000 Transcriptional regulation of adipogenesis. Genes and Development 14:1293-1307 29. Amri EZ, Dani C, Doglio A, Etienne J, Grimaldi P, Ailhaud G 1986 Adipose cell differentiation: evidence for a two-step process in the polyamine-dependent Ob1754 clonal line. Biochem J 238:115-122 30. Schoonjans K, Staels B, Auwerx J 1996 The peroxisome proliferator activated receptors (PPARS) and their effects on lipid metabolism and adipocyte differentiation. Biochim Biophys Acta 1302:93-109 31. Elberg G, Gimble JM, Tsai SY 2000 Modulation of the murine peroxisome proliferator- activated receptor gamma 2 promoter activity by CCAAT/enhancer-binding proteins. Journal of Biological Chemistry 275:27815-27822 32. Yeh WC, Cao Z, Classon M, McKnight SL 1995 Cascade regulation of terminal adipocyte differentiation by three members of the C/EBP family of leucine zipper proteins. Genes and Development 9:168-181 33. Tanaka T, Yoshida N, Kishimoto T, Akira S 1997 Defective adipocyte differentiation in mice lacking the C/EBPbeta and/or C/EBPdelta gene. EMBO J 16:7432-7443 34. Zhu Y, Qi C, Korenberg JR, Chen XN, Noya D, Rao MS, Reddy JK 1995 Structural organization of mouse peroxisome proliferator-activated receptor gamma (mPPAR gamma) gene: alternative promoter use and different splicing yield two mPPAR gamma isoforms. Proc Natl Acad Sci U S A 92:7921-7925 35. Ren D, Collingwood TN, Rebar EJ, Wolffe AP, Camp HS 2002 PPARgamma knockdown by engineered transcription factors: exogenous PPARgamma2 but not PPARgamma1 reactivates adipogenesis. Genes and Development 16:27-32 36. Rosen ED, Sarraf P, Troy AE, Bradwin G, Moore K, Milstone DS, Spiegelman BM, Mortensen RM 1999 PPAR gamma is required for the differentiation of adipose tissue in vivo and in vitro. Mol Cell 4:611-617 37. Wang ND, Finegold MJ, Bradley A, Ou CN, Abdelsayed SV, Wilde MD, Taylor LR, Wilson DR, Darlington GJ 1995 Impaired energy homeostasis in C/EBP alpha knockout mice. Science 269:1108-1112

90 38. Wu Z, Rosen ED, Brun R, Hauser S, Adelmant G, Troy AE, McKeon C, Darlington GJ, Spiegelman BM 1999 Cross-regulation of C/EBP alpha and PPAR gamma controls the transcriptional pathway of adipogenesis and insulin sensitivity. Mol Cell 3:151-158 39. Rosen ED, Hsu CH, Wang X, Sakai S, Freeman MW, Gonzalez FJ, Spiegelman BM 2002 C/EBPalpha induces adipogenesis through PPARgamma: a unified pathway. Genes and Development 16:22-26 40. Sul HS, Smas C, Mei B, Zhou L 2000 Function of pref-1 as an inhibitor of adipocyte differentiation. Int J Obes Relat Metab Disord 24 Suppl 4:S15-S19 41. Kuri-Harcuch W, Wise LS, Green H 1978 Interruption of the adipose conversion of 3T3 cells by biotin deficiency: differentiation without triglyceride accumulation. Cell 14:53- 59 42. Garcia dH, Birnbaum MJ 1989 The regulation by insulin of glucose transporter gene expression in 3T3 adipocytes. Journal of Biological Chemistry 264:9885-9890 43. Guest SJ, Hadcock JR, Watkins DC, Malbon CC 1990 Beta 1- and beta 2-adrenergic receptor expression in differentiating 3T3-L1 cells. Independent regulation at the level of mRNA. Journal of Biological Chemistry 265:5370-5375 44. Spiegelman BM, Frank M, Green H 1983 Molecular cloning of mRNA from 3T3 adipocytes. Regulation of mRNA content for glycerophosphate dehydrogenase and other differentiation-dependent proteins during adipocyte development. Journal of Biological Chemistry 258:10083-10089 45. MacDougald OA, Hwang CS, Fan H, Lane MD 1995 Regulated expression of the obese gene product (leptin) in white adipose tissue and 3T3-L1 adipocytes. Proc Natl Acad Sci U S A 92:9034-9037 46. Hollenberg AN, Susulic VS, Madura JP, Zhang B, Moller DE, Tontonoz P, Sarraf P, Spiegelman BM, Lowell BB 1997 Functional antagonism between CCAAT/Enhancer binding protein-alpha and peroxisome proliferator-activated receptor-gamma on the leptin promoter. Journal of Biological Chemistry 272:5283-5290 47. Harp JB 2004 New insights into inhibitors of adipogenesis. Curr Opin Lipidol 15:303- 307 48. Goldberg IJ 1996 Lipoprotein lipase and lipolysis: central roles in lipoprotein metabolism and atherogenesis. J Lipid Res 37:693-707 49. Brunzell JD, Hazzard WR, Porte D, Jr., Bierman EL 1973 Evidence for a common, saturable, triglyceride removal mechanism for chylomicrons and very low density lipoproteins in man. J Clin Invest 52:1578-1585 50. Gruffat D, Durand D, Graulet B, Bauchart D 1996 Regulation of VLDL synthesis and secretion in the liver. Reprod Nutr Dev 36:375-389 51. Williams CM, Bateman PA, Jackson KG, Yaqoob P 2004 Dietary fatty acids and chylomicron synthesis and secretion. Biochemal Society Transections 32:55-58 52. Merkel M, Heeren J, Dudeck W, Rinninger F, Radner H, Breslow JL, Goldberg IJ, Zechner R, Greten H 2002 Inactive lipoprotein lipase (LPL) alone increases selective cholesterol ester uptake in vivo, whereas in the presence of active LPL it also increases triglyceride hydrolysis and whole particle lipoprotein uptake. Journal of Biological Chemistry 277:7405-7411 53. Medh JD, Bowen SL, Fry GL, Ruben S, Andracki M, Inoue I, Lalouel JM, Strickland DK, Chappell DA 1996 Lipoprotein lipase binds to low density lipoprotein receptors and

91 induces receptor-mediated catabolism of very low density lipoproteins in vitro. Journal of Biological Chemistry 271:17073-17080 54. Seo T, Al Haideri M, Treskova E, Worgall TS, Kako Y, Goldberg IJ, Deckelbaum RJ 2000 Lipoprotein lipase-mediated selective uptake from low density lipoprotein requires cell surface proteoglycans and is independent of scavenger receptor class B type 1. Journal of Biological Chemistry 275:30355-30362 55. Yacoub LK, Vanni TM, Goldberg IJ 1990 Lipoprotein lipase mRNA in neonatal and adult mouse tissues: comparison of normal and combined lipase deficiency (cld) mice assessed by in situ hybridization. J Lipid Res 31:1845-1852 56. Semenkovich CF, Chen SH, Wims M, Luo CC, Li WH, Chan L 1989 Lipoprotein lipase and hepatic lipase mRNA tissue specific expression, developmental regulation, and evolution. J Lipid Res 30:423-431 57. Augustus A, Yagyu H, Haemmerle G, Bensadoun A, Vikramadithyan RK, Park SY, Kim JK, Zechner R, Goldberg IJ 2004 Cardiac-specific knock-out of lipoprotein lipase alters plasma lipoprotein triglyceride metabolism and cardiac gene expression. Journal of Biological Chemistry 279:25050-25057 58. Kim JK, Fillmore JJ, Chen Y, Yu C, Moore IK, Pypaert M, Lutz EP, Kako Y, Velez- Carrasco W, Goldberg IJ, Breslow JL, Shulman GI 2001 Tissue-specific overexpression of lipoprotein lipase causes tissue-specific insulin resistance. Proc Natl Acad Sci U S A 98:7522-7527 59. Wong H, Schotz MC 2002 The lipase gene family. J Lipid Res 43:993-999 60. Merkel M, Eckel RH, Goldberg IJ 2002 Lipoprotein lipase: genetics, lipid uptake, and regulation. J Lipid Res 43:1997-2006 61. Semenkovich CF, Wims M, Noe L, Etienne J, Chan L 1989 Insulin regulation of lipoprotein lipase activity in 3T3-L1 adipocytes is mediated at posttranscriptional and posttranslational levels. Journal of Biological Chemistry 264:9030-9038 62. Hua XX, Enerback S, Hudson J, Youkhana K, Gimble JM 1991 Cloning and characterization of the promoter of the murine lipoprotein lipase-encoding gene: structural and functional analysis. Gene 107:247-258 63. Schoonjans K, Peinado-Onsurbe J, Lefebvre AM, Heyman RA, Briggs M, Deeb S, Staels B, Auwerx J 1996 PPARalpha and PPARgamma activators direct a distinct tissue- specific transcriptional response via a PPRE in the lipoprotein lipase gene. EMBO J 15:5336-5348 64. Kim JB, Spiegelman BM 1996 ADD1/SREBP1 promotes adipocyte differentiation and gene expression linked to fatty acid metabolism. Genes and Development 10:1096-1107 65. Hogan JC, Stephens JM 2003 STAT 1 binds to the LPL promoter in vitro. Biochem Biophys Res Commun 307:350-354 66. Homma H, Kurachi H, Nishio Y, Takeda T, Yamamoto T, Adachi K, Morishige K, Ohmichi M, Matsuzawa Y, Murata Y 2000 Estrogen suppresses transcription of lipoprotein lipase gene. Existence of a unique estrogen response element on the lipoprotein lipase promoter. Journal of Biological Chemistry 275:11404-11411 67. Morin CL, Schlaepfer IR, Eckel RH 1995 Tumor necrosis factor-alpha eliminates binding of NF-Y and an octamer-binding protein to the lipoprotein lipase promoter in 3T3-L1 adipocytes. J Clin Invest 95:1684-1689 68. Ranganathan G, Phan D, Pokrovskaya ID, McEwen JE, Li C, Kern PA 2002 The translational regulation of lipoprotein lipase by epinephrine involves an RNA binding

92 complex including the catalytic subunit of protein kinase A. Journal of Biological Chemistry 277:43281-43287 69. Jenni S, Leibundgut M, Maier T, Ban N 2006 Architecture of a fungal fatty acid synthase at 5 A resolution. Science 311:1263-1267 70. Maier T, Jenni S, Ban N 2006 Architecture of mammalian fatty acid synthase at 4.5 A resolution. Science 311:1258-1262 71. Paulauskis JD, Sul HS 1989 Hormonal regulation of mouse fatty acid synthase gene transcription in liver. Journal of Biological Chemistry 264:574-577 72. Hillgartner FB, Salati LM, Goodridge AG 1995 Physiological and molecular mechanisms involved in nutritional regulation of fatty acid synthesis. Physiol Rev 75:47-76 73. Sul HS, Latasa MJ, Moon Y, Kim KH 2000 Regulation of the fatty acid synthase promoter by insulin. J Nutr 130:315S-320S 74. Stapleton SR, Mitchell DA, Salati LM, Goodridge AG 1990 Triiodothyronine stimulates transcription of the fatty acid synthase gene in chick embryo hepatocytes in culture. Insulin and insulin-like growth factor amplify that effect. Journal of Biological Chemistry 265:18442-18446 75. Lakshmanan MR, Nepokroeff CM, Porter JW 1972 Control of the synthesis of fatty-acid synthetase in rat liver by insulin, glucagon, and adenosine 3':5' cyclic monophosphate. Proc Natl Acad Sci U S A 69:3516-3519 76. Semenkovich CF 1997 Regulation of fatty acid synthase (FAS). Prog Lipid Res 36:43-53 77. Moon YS, Latasa MJ, Kim KH, Wang D, Sul HS 2000 Two 5'-regions are required for nutritional and insulin regulation of the fatty-acid synthase promoter in transgenic mice. Journal of Biological Chemistry 275:10121-10127 78. Shimano H, Yahagi N, Amemiya-Kudo M, Hasty AH, Osuga J, Tamura Y, Shionoiri F, Iizuka Y, Ohashi K, Harada K, Gotoda T, Ishibashi S, Yamada N 1999 Sterol regulatory element-binding protein-1 as a key transcription factor for nutritional induction of lipogenic enzyme genes. Journal of Biological Chemistry 274:35832-35839 79. Hollenberg CH, RABEN MS, ASTWOOD EB 1961 The lipolytic response to corticotropin. Endocrinology 68:589-598 80. VAUGHAN M, BERGER JE, STEINBERG D 1964 HORMONE-SENSITIVE LIPASE AND MONOGLYCERIDE LIPASE ACTIVITIES IN ADIPOSE TISSUE. Journal of Biological Chemistry 239:401-409 81. Li Z, Sumida M, Birchbauer A, Schotz MC, Reue K 1994 Isolation and characterization of the gene for mouse hormone-sensitive lipase. Genomics 24:259-265 82. Grober J, Laurell H, Blaise R, Fabry B, Schaak S, Holm C, Langin D 1997 Characterization of the promoter of human adipocyte hormone-sensitive lipase. Biochem J 328 ( Pt 2):453-461 83. Mairal A, Melaine N, Laurell H, Grober J, Holst LS, Guillaudeux T, Holm C, Jegou B, Langin D 2002 Characterization of a novel testicular form of human hormone-sensitive lipase. Biochem Biophys Res Commun 291:286-290 84. Laurin NN, Wang SP, Mitchell GA 2000 The hormone-sensitive lipase gene is transcribed from at least five alternative first exons in mouse adipose tissue. Mamm Genome 11:972-978 85. Mulder H, Holst LS, Svensson H, Degerman E, Sundler F, Ahren B, Rorsman P, Holm C 1999 Hormone-sensitive lipase, the rate-limiting enzyme in triglyceride hydrolysis, is expressed and active in beta-cells. Diabetes 48:228-232

93 86. Holm C, Kirchgessner TG, Svenson KL, Lusis AJ, Belfrage P, Schotz MC 1988 Nucleotide sequence of rat adipose hormone sensitive lipase cDNA. Nucleic Acids Res 16:9879 87. Osterlund T, Danielsson B, Degerman E, Contreras JA, Edgren G, Davis RC, Schotz MC, Holm C 1996 Domain-structure analysis of recombinant rat hormone-sensitive lipase. Biochem J 319 ( Pt 2):411-420 88. Shen WJ, Patel S, Hong R, Kraemer FB 2000 Hormone-sensitive lipase functions as an oligomer. Biochemistry 39:2392-2398 89. Fredrikson G, Stralfors P, Nilsson NO, Belfrage P 1981 Hormone-sensitive lipase of rat adipose tissue. Purification and some properties. Journal of Biological Chemistry 256:6311-6320 90. Fredrikson G, Tornqvist H, Belfrage P 1986 Hormone-sensitive lipase and monoacylglycerol lipase are both required for complete degradation of adipocyte triacylglycerol. Biochim Biophys Acta 876:288-293 91. Fredrikson G, Belfrage P 1983 Positional specificity of hormone-sensitive lipase from rat adipose tissue. Journal of Biological Chemistry 258:14253-14256 92. Anthonsen MW, Ronnstrand L, Wernstedt C, Degerman E, Holm C 1998 Identification of novel phosphorylation sites in hormone-sensitive lipase that are phosphorylated in response to isoproterenol and govern activation properties in vitro. Journal of Biological Chemistry 273:215-221 93. Greenberg AS, Shen WJ, Muliro K, Patel S, Souza SC, Roth RA, Kraemer FB 2001 Stimulation of lipolysis and hormone-sensitive lipase via the extracellular signal- regulated kinase pathway. Journal of Biological Chemistry 276:45456-45461 94. Garton AJ, Campbell DG, Carling D, Hardie DG, Colbran RJ, Yeaman SJ 1989 Phosphorylation of bovine hormone-sensitive lipase by the AMP-activated protein kinase. A possible antilipolytic mechanism. Eur J Biochem 179:249-254 95. Clifford GM, Kraemer FB, Yeaman SJ, Vernon RG 2001 Translocation of hormone- sensitive lipase and perilipin upon lipolytic stimulation during the lactation cycle of the rat. Metabolism 50:1264-1269 96. Sztalryd C, Kraemer FB 1994 Differences in hormone-sensitive lipase expression in white adipose tissue from various anatomic locations of the rat. Metabolism 43:241-247 97. Fortier M, Soni K, Laurin N, Wang SP, Mauriege P, Jirik FR, Mitchell GA 2005 Human hormone-sensitive lipase (HSL): expression in white fat corrects the white adipose phenotype of HSL-deficient mice. J Lipid Res 46:1860-1867 98. Osuga J, Ishibashi S, Oka T, Yagyu H, Tozawa R, Fujimoto A, Shionoiri F, Yahagi N, Kraemer FB, Tsutsumi O, Yamada N 2000 Targeted disruption of hormone-sensitive lipase results in male sterility and adipocyte hypertrophy, but not in obesity. Proc Natl Acad Sci U S A 97:787-792 99. Egan JJ, Greenberg AS, Chang MK, Wek SA, Moos MC, Jr., Londos C 1992 Mechanism of hormone-stimulated lipolysis in adipocytes: translocation of hormone-sensitive lipase to the lipid storage droplet. Proc Natl Acad Sci U S A 89:8537-8541 100. Brasaemle DL, Levin DM, Adler-Wailes DC, Londos C 2000 The lipolytic stimulation of 3T3-L1 adipocytes promotes the translocation of hormone-sensitive lipase to the surfaces of lipid storage droplets. Biochim Biophys Acta 1483:251-262

94 101. Sztalryd C, Xu G, Dorward H, Tansey JT, Contreras JA, Kimmel AR, Londos C 2003 Perilipin A is essential for the translocation of hormone-sensitive lipase during lipolytic activation. Journal of Cell Biology 161:1093-1103 102. Servetnick DA, Brasaemle DL, Gruia-Gray J, Kimmel AR, Wolff J, Londos C 1995 Perilipins are associated with cholesteryl ester droplets in steroidogenic adrenal cortical and Leydig cells. Journal of Biological Chemistry 270:16970-16973 103. Greenberg AS, Egan JJ, Wek SA, Moos MC, Jr., Londos C, Kimmel AR 1993 Isolation of cDNAs for perilipins A and B: sequence and expression of lipid droplet-associated proteins of adipocytes. Proc Natl Acad Sci U S A 90:12035-12039 104. Greenberg AS, Egan JJ, Wek SA, Garty NB, Blanchette-Mackie EJ, Londos C 1991 Perilipin, a major hormonally regulated adipocyte-specific phosphoprotein associated with the periphery of lipid storage droplets. Journal of Biological Chemistry 266:11341- 11346 105. Souza SC, de Vargas LM, Yamamoto MT, Lien P, Franciosa MD, Moss LG, Greenberg AS 1998 Overexpression of perilipin A and B blocks the ability of tumor necrosis factor alpha to increase lipolysis in 3T3-L1 adipocytes. Journal of Biological Chemistry 273:24665-24669 106. Tansey JT, Huml AM, Vogt R, Davis KE, Jones JM, Fraser KA, Brasaemle DL, Kimmel AR, Londos C 2003 Functional studies on native and mutated forms of perilipins. A role in protein kinase A-mediated lipolysis of triacylglycerols. Journal of Biological Chemistry 278:8401-8406 107. Tansey JT, Sztalryd C, Gruia-Gray J, Roush DL, Zee JV, Gavrilova O, Reitman ML, Deng CX, Li C, Kimmel AR, Londos C 2001 Perilipin ablation results in a lean mouse with aberrant adipocyte lipolysis, enhanced leptin production, and resistance to diet- induced obesity. Proc Natl Acad Sci U S A 98:6494-6499 108. Miyoshi H, Souza SC, Zhang HH, Strissel KJ, Christoffolete MA, Kovsan J, Rudich A, Kraemer FB, Bianco AC, Obin MS, Greenberg AS 2006 Perilipin promotes hormone- sensitive lipase-mediated adipocyte lipolysis via phosphorylation-dependent and - independent mechanisms. Journal of Biological Chemistry 281:15837-15844 109. Robenek H, Robenek MJ, Buers I, Lorkowski S, Hofnagel O, Troyer D, Severs NJ 2005 Lipid droplets gain PAT family proteins by interaction with specialized plasma membrane domains. Journal of Biological Chemistry 280:26330-26338 110. Aboulaich N, Vener AV, Stralfors P 2006 Hormonal control of reversible translocation of perilipin B to the plasma membrane in primary human adipocytes. Journal of Biological Chemistry 281:11446-11449 111. Okazaki H, Osuga J, Tamura Y, Yahagi N, Tomita S, Shionoiri F, Iizuka Y, Ohashi K, Harada K, Kimura S, Gotoda T, Shimano H, Yamada N, Ishibashi S 2002 Lipolysis in the absence of hormone-sensitive lipase: evidence for a common mechanism regulating distinct lipases. Diabetes 51:3368-3375 112. Zimmermann R, Strauss JG, Haemmerle G, Schoiswohl G, Birner-Gruenberger R, Riederer M, Lass A, Neuberger G, Eisenhaber F, Hermetter A, Zechner R 2004 Fat mobilization in adipose tissue is promoted by adipose triglyceride lipase. Science 306:1383-1386 113. Villena JA, Roy S, Sarkadi-Nagy E, Kim KH, Sul HS 2004 Desnutrin, an adipocyte gene encoding a novel patatin domain-containing protein, is induced by fasting and

95 glucocorticoids: ectopic expression of desnutrin increases triglyceride hydrolysis. Journal of Biological Chemistry 279:47066-47075 114. Kershaw EE, Flier JS 2004 Adipose tissue as an endocrine organ. J Clin Endocrinol Metab 89:2548-2556 115. Wolf G 2005 The mechanism and regulation of fat mobilization from adipose tissue: desnutrin, a newly discovered lipolytic enzyme. Nutr Rev 63:166-170 116. Joost HG, Bell GI, Best JD, Birnbaum MJ, Charron MJ, Chen YT, Doege H, James DE, Lodish HF, Moley KH, Moley JF, Mueckler M, Rogers S, Schurmann A, Seino S, Thorens B 2002 Nomenclature of the GLUT/SLC2A family of sugar/polyol transport facilitators. Am J Physiol Endocrinol Metab 282:E974-E976 117. Joost HG, Thorens B 2001 The extended GLUT-family of sugar/polyol transport facilitators: nomenclature, sequence characteristics, and potential function of its novel members (review). Mol Membr Biol 18:247-256 118. Wood IS, Trayhurn P 2003 Glucose transporters (GLUT and SGLT): expanded families of sugar transport proteins. Br J Nutr 89:3-9 119. Malide D, Ramm G, Cushman SW, Slot JW 2000 Immunoelectron microscopic evidence that GLUT4 translocation explains the stimulation of glucose transport in isolated rat white adipose cells. J Cell Sci 113 Pt 23:4203-4210 120. Ploug T, Ralston E 2002 Exploring the whereabouts of GLUT4 in skeletal muscle (review). Mol Membr Biol 19:39-49 121. Karylowski O, Zeigerer A, Cohen A, McGraw TE 2004 GLUT4 is retained by an intracellular cycle of vesicle formation and fusion with endosomes. Mol Biol Cell 15:870-882 122. Govers R, Coster AC, James DE 2004 Insulin increases cell surface GLUT4 levels by dose dependently discharging GLUT4 into a cell surface recycling pathway. Mol Cell Biol 24:6456-6466 123. Watson RT, Pessin JE 2006 Bridging the GAP between insulin signaling and GLUT4 translocation. Trends Biochem Sci 31:215-222 124. Martin OJ, Lee A, McGraw TE 2006 GLUT4 distribution between the plasma membrane and the intracellular compartments is maintained by an insulin-modulated bipartite dynamic mechanism. Journal of Biological Chemistry 281:484-490 125. el Jack AK, Kandror KV, Pilch PF 1999 The formation of an insulin-responsive vesicular cargo compartment is an early event in 3T3-L1 adipocyte differentiation. Mol Biol Cell 10:1581-1594 126. Watson RT, Kanzaki M, Pessin JE 2004 Regulated membrane trafficking of the insulin- responsive glucose transporter 4 in adipocytes. Endocr Rev 25:177-204 127. White MF 2002 IRS proteins and the common path to diabetes. Am J Physiol Endocrinol Metab 283:E413-E422 128. Watson RT, Pessin JE 2001 Intracellular organization of insulin signaling and GLUT4 translocation. Recent Prog Horm Res 56:175-193 129. Baumann CA, Ribon V, Kanzaki M, Thurmond DC, Mora S, Shigematsu S, Bickel PE, Pessin JE, Saltiel AR 2000 CAP defines a second signalling pathway required for insulin- stimulated glucose transport. Nature 407:202-207 130. Liu J, Kimura A, Baumann CA, Saltiel AR 2002 APS facilitates c-Cbl tyrosine phosphorylation and GLUT4 translocation in response to insulin in 3T3-L1 adipocytes. Mol Cell Biol 22:3599-3609

96 131. Ahmed Z, Pillay TS 2003 Adapter protein with a pleckstrin homology (PH) and an Src homology 2 (SH2) domain (APS) and SH2-B enhance insulin-receptor autophosphorylation, extracellular-signal-regulated kinase and phosphoinositide 3- kinase-dependent signalling. Biochem J 371:405-412 132. Kitamura T, Kahn CR, Accili D 2003 Insulin receptor knockout mice. Annu Rev Physiol 65:313-332 133. Liu SC, Wang Q, Lienhard GE, Keller SR 1999 Insulin receptor substrate 3 is not essential for growth or glucose homeostasis. Journal of Biological Chemistry 274:18093- 18099 134. Fantin VR, Wang Q, Lienhard GE, Keller SR 2000 Mice lacking insulin receptor substrate 4 exhibit mild defects in growth, reproduction, and glucose homeostasis. Am J Physiol Endocrinol Metab 278:E127-E133 135. Nishizawa Y, Bray GA 1980 Evidence for a circulating ergostatic factor: studies on parabiotic rats. Am J Physiol 239:R344-R351 136. Cook KS, Min HY, Johnson D, Chaplinsky RJ, Flier JS, Hunt CR, Spiegelman BM 1987 Adipsin: a circulating serine protease homolog secreted by adipose tissue and sciatic nerve. Science 237:402-405 137. Zhang Y, Proenca R, Maffei M, Barone M, Leopold L, Friedman JM 1994 Positional cloning of the mouse obese gene and its human homologue. Nature 372:425-432 138. Kline AD, Becker GW, Churgay LM, Landen BE, Martin DK, Muth WL, Rathnachalam R, Richardson JM, Schoner B, Ulmer M, Hale JE 1997 Leptin is a four-helix bundle: secondary structure by NMR. FEBS Lett 407:239-242 139. Pelleymounter MA, Cullen MJ, Baker MB, Hecht R, Winters D, Boone T, Collins F 1995 Effects of the obese gene product on body weight regulation in ob/ob mice. Science 269:540-543 140. Campfield LA, Smith FJ, Guisez Y, Devos R, Burn P 1995 Recombinant mouse OB protein: evidence for a peripheral signal linking adiposity and central neural networks. Science 269:546-549 141. Tartaglia LA, Dembski M, Weng X, Deng N, Culpepper J, Devos R, Richards GJ, Campfield LA, Clark FT, Deeds J, Muir C, Sanker S, Moriarty A, Moore KJ, Smutko JS, Mays GG, Wool EA, Monroe CA, Tepper RI 1995 Identification and expression cloning of a leptin receptor, OB-R. Cell 83:1263-1271 142. Lee GH, Proenca R, Montez JM, Carroll KM, Darvishzadeh JG, Lee JI, Friedman JM 1996 Abnormal splicing of the leptin receptor in diabetic mice. Nature 379:632-635 143. Barr VA, Lane K, Taylor SI 1999 Subcellular localization and internalization of the four human leptin receptor isoforms. Journal of Biological Chemistry 274:21416-21424 144. Ghilardi N, Ziegler S, Wiestner A, Stoffel R, Heim MH, Skoda RC 1996 Defective STAT signaling by the leptin receptor in diabetic mice. Proc Natl Acad Sci U S A 93:6231-6235 145. Moschos S, Chan JL, Mantzoros CS 2002 Leptin and reproduction: a review. Fertil Steril 77:433-444 146. Dal Farra C, Zsurger N, Vincent JP, Cupo A 2000 Binding of a pure 125I- monoiodoleptin analog to mouse tissues: a developmental study. Peptides 21:577-587 147. Huang L, Wang Z, Li C 2001 Modulation of circulating leptin levels by its soluble receptor. Journal of Biological Chemistry 276:6343-6349

97 148. Considine RV, Sinha MK, Heiman ML, Kriauciunas A, Stephens TW, Nyce MR, Ohannesian JP, Marco CC, McKee LJ, Bauer TL, . 1996 Serum immunoreactive-leptin concentrations in normal-weight and obese humans. N Engl J Med 334:292-295 149. Frederich RC, Hamann A, Anderson S, Lollmann B, Lowell BB, Flier JS 1995 Leptin levels reflect body lipid content in mice: evidence for diet-induced resistance to leptin action. Nat Med 1:1311-1314 150. Hamilton BS, Paglia D, Kwan AY, Deitel M 1995 Increased obese mRNA expression in omental fat cells from massively obese humans. Nat Med 1:953-956 151. Montague CT, Prins JB, Sanders L, Digby JE, O'Rahilly S 1997 Depot- and sex-specific differences in human leptin mRNA expression: implications for the control of regional fat distribution. Diabetes 46:342-347 152. Havel PJ, Kasim-Karakas S, Dubuc GR, Mueller W, Phinney SD 1996 Gender differences in plasma leptin concentrations. Nat Med 2:949-950 153. Landt M, Gingerich RL, Havel PJ, Mueller WM, Schoner B, Hale JE, Heiman ML 1998 Radioimmunoassay of rat leptin: sexual dimorphism reversed from humans. Clin Chem 44:565-570 154. Weigle DS, Duell PB, Connor WE, Steiner RA, Soules MR, Kuijper JL 1997 Effect of fasting, refeeding, and dietary fat restriction on plasma leptin levels. J Clin Endocrinol Metab 82:561-565 155. Havel PJ 1999 Mechanisms regulating leptin production: implications for control of energy balance. Am J Clin Nutr 70:305-306 156. Friedman JM 1998 Leptin, leptin receptors, and the control of body weight. Nutr Rev 56:s38-s46 157. Ahima RS, Prabakaran D, Mantzoros C, Qu D, Lowell B, Maratos-Flier E, Flier JS 1996 Role of leptin in the neuroendocrine response to fasting. Nature 382:250-252 158. Caro JF, Sinha MK, Kolaczynski JW, Zhang PL, Considine RV 1996 Leptin: the tale of an obesity gene. Diabetes 45:1455-1462 159. Flier JS 2004 Obesity wars: molecular progress confronts an expanding epidemic. Cell 116:337-350 160. Bornstein SR, Uhlmann K, Haidan A, Ehrhart-Bornstein M, Scherbaum WA 1997 Evidence for a novel peripheral action of leptin as a metabolic signal to the adrenal gland: leptin inhibits cortisol release directly. Diabetes 46:1235-1238 161. Cervero A, Dominguez F, Horcajadas JA, Quinonero A, Pellicer A, Simon C 2006 The role of the leptin in reproduction. Curr Opin Obstet Gynecol 18:297-303 162. Ahima RS, Saper CB, Flier JS, Elmquist JK 2000 Leptin regulation of neuroendocrine systems. Front Neuroendocrinol 21:263-307 163. Baskin DG, Hahn TM, Schwartz MW 1999 Leptin sensitive neurons in the hypothalamus. Horm Metab Res 31:345-350 164. Jequier E 2002 Leptin signaling, adiposity, and energy balance. Ann N Y Acad Sci 967:379-388 165. Schwartz MW, Seeley RJ, Campfield LA, Burn P, Baskin DG 1996 Identification of targets of leptin action in rat hypothalamus. J Clin Invest 98:1101-1106 166. Simerly RB 2005 Wired on hormones: endocrine regulation of hypothalamic development. Curr Opin Neurobiol 15:81-85 167. Seufert J 2004 Leptin effects on pancreatic beta-cell gene expression and function. Diabetes 53 Suppl 1:S152-S158

98 168. Havel PJ 2000 Role of adipose tissue in body-weight regulation: mechanisms regulating leptin production and energy balance. Proc Nutr Soc 59:359-371 169. Scherer PE, Williams S, Fogliano M, Baldini G, Lodish HF 1995 A novel serum protein similar to C1q, produced exclusively in adipocytes. Journal of Biological Chemistry 270:26746-26749 170. Maeda K, Okubo K, Shimomura I, Funahashi T, Matsuzawa Y, Matsubara K 1996 cDNA cloning and expression of a novel adipose specific collagen-like factor, apM1 (AdiPose Most abundant Gene transcript 1). Biochem Biophys Res Commun 221:286-289 171. Hu E, Liang P, Spiegelman BM 1996 AdipoQ is a novel adipose-specific gene dysregulated in obesity. Journal of Biological Chemistry 271:10697-10703 172. Nakano Y, Tobe T, Choi-Miura NH, Mazda T, Tomita M 1996 Isolation and characterization of GBP28, a novel gelatin-binding protein purified from human plasma. J Biochem (Tokyo) 120:803-812 173. Das K, Lin Y, Widen E, Zhang Y, Scherer PE 2001 Chromosomal localization, expression pattern, and promoter analysis of the mouse gene encoding adipocyte-specific secretory protein Acrp30. Biochem Biophys Res Commun 280:1120-1129 174. Berg AH, Combs TP, Scherer PE 2002 ACRP30/adiponectin: an adipokine regulating glucose and lipid metabolism. Trends Endocrinol Metab 13:84-89 175. Shapiro L, Scherer PE 1998 The crystal structure of a complement-1q family protein suggests an evolutionary link to tumor necrosis factor. Curr Biol 8:335-338 176. Pajvani UB, Du X, Combs TP, Berg AH, Rajala MW, Schulthess T, Engel J, Brownlee M, Scherer PE 2003 Structure-function studies of the adipocyte-secreted hormone Acrp30/adiponectin. Implications fpr metabolic regulation and bioactivity. Journal of Biological Chemistry 278:9073-9085 177. Waki H, Yamauchi T, Kamon J, Ito Y, Uchida S, Kita S, Hara K, Hada Y, Vasseur F, Froguel P, Kimura S, Nagai R, Kadowaki T 2003 Impaired multimerization of human adiponectin mutants associated with diabetes. Molecular structure and multimer formation of adiponectin. Journal of Biological Chemistry 278:40352-40363 178. Fruebis J, Tsao TS, Javorschi S, Ebbets-Reed D, Erickson MR, Yen FT, Bihain BE, Lodish HF 2001 Proteolytic cleavage product of 30-kDa adipocyte complement-related protein increases fatty acid oxidation in muscle and causes weight loss in mice. Proc Natl Acad Sci U S A 98:2005-2010 179. Wang Y, Xu A, Knight C, Xu LY, Cooper GJ 2002 Hydroxylation and glycosylation of the four conserved lysine residues in the collagenous domain of adiponectin. Potential role in the modulation of its insulin-sensitizing activity. Journal of Biological Chemistry 277:19521-19529 180. Bottner A, Kratzsch J, Muller G, Kapellen TM, Bluher S, Keller E, Bluher M, Kiess W 2004 Gender differences of adiponectin levels develop during the progression of puberty and are related to serum androgen levels. J Clin Endocrinol Metab 89:4053-4061 181. Yamauchi T, Kamon J, Ito Y, Tsuchida A, Yokomizo T, Kita S, Sugiyama T, Miyagishi M, Hara K, Tsunoda M, Murakami K, Ohteki T, Uchida S, Takekawa S, Waki H, Tsuno NH, Shibata Y, Terauchi Y, Froguel P, Tobe K, Koyasu S, Taira K, Kitamura T, Shimizu T, Nagai R, Kadowaki T 2003 Cloning of adiponectin receptors that mediate antidiabetic metabolic effects. Nature 423:762-769 182. Nilsson L, Binart N, Bohlooly Y, Bramnert M, Egecioglu E, Kindblom J, Kelly PA, Kopchick JJ, Ormandy CJ, Ling C, Billig H 2005 Prolactin and growth hormone regulate

99 adiponectin secretion and receptor expression in adipose tissue. Biochem Biophys Res Commun 331:1120-1126 183. Hardie DG, Carling D 1997 The AMP-activated protein kinase--fuel gauge of the mammalian cell? Eur J Biochem 246:259-273 184. Gil-Campos M, Canete RR, Gil A 2004 Adiponectin, the missing link in insulin resistance and obesity. Clin Nutr 23:963-974 185. Kharroubi I, Rasschaert J, Eizirik DL, Cnop M 2003 Expression of adiponectin receptors in pancreatic beta cells. Biochem Biophys Res Commun 312:1118-1122 186. Matsuzawa Y, Funahashi T, Kihara S, Shimomura I 2004 Adiponectin and metabolic syndrome. Arterioscler Thromb Vasc Biol 24:29-33 187. Yamamoto Y, Hirose H, Saito I, Nishikai K, Saruta T 2004 Adiponectin, an adipocyte- derived protein, predicts future insulin resistance: two-year follow-up study in Japanese population. J Clin Endocrinol Metab 89:87-90 188. Spranger J, Kroke A, Mohlig M, Bergmann MM, Ristow M, Boeing H, Pfeiffer AF 2003 Adiponectin and protection against type 2 diabetes mellitus. Lancet 361:226-228 189. Lindsay RS, Funahashi T, Hanson RL, Matsuzawa Y, Tanaka S, Tataranni PA, Knowler WC, Krakoff J 2002 Adiponectin and development of type 2 diabetes in the Pima Indian population. Lancet 360:57-58 190. Kubota N, Terauchi Y, Yamauchi T, Kubota T, Moroi M, Matsui J, Eto K, Yamashita T, Kamon J, Satoh H, Yano W, Froguel P, Nagai R, Kimura S, Kadowaki T, Noda T 2002 Disruption of adiponectin causes insulin resistance and neointimal formation. Journal of Biological Chemistry 277:25863-25866 191. Combs TP, Pajvani UB, Berg AH, Lin Y, Jelicks LA, Laplante M, Nawrocki AR, Rajala MW, Parlow AF, Cheeseboro L, Ding YY, Russell RG, Lindemann D, Hartley A, Baker GR, Obici S, Deshaies Y, Ludgate M, Rossetti L, Scherer PE 2004 A transgenic mouse with a deletion in the collagenous domain of adiponectin displays elevated circulating adiponectin and improved insulin sensitivity. Endocrinology 145:367-383 192. Berg AH, Combs TP, Du X, Brownlee M, Scherer PE 2001 The adipocyte-secreted protein Acrp30 enhances hepatic insulin action. Nat Med 7:947-953 193. Duntas LH, Popovic V, Panotopoulos G 2004 Adiponectin: novelties in metabolism and hormonal regulation. Nutr Neurosci 7:195-200 194. Qi Y, Takahashi N, Hileman SM, Patel HR, Berg AH, Pajvani UB, Scherer PE, Ahima RS 2004 Adiponectin acts in the brain to decrease body weight. Nat Med 10:524-529 195. Hotta K, Funahashi T, Arita Y, Takahashi M, Matsuda M, Okamoto Y, Iwahashi H, Kuriyama H, Ouchi N, Maeda K, Nishida M, Kihara S, Sakai N, Nakajima T, Hasegawa K, Muraguchi M, Ohmoto Y, Nakamura T, Yamashita S, Hanafusa T, Matsuzawa Y 2000 Plasma concentrations of a novel, adipose-specific protein, adiponectin, in type 2 diabetic patients. Arterioscler Thromb Vasc Biol 20:1595-1599 196. Okamoto Y, Arita Y, Nishida M, Muraguchi M, Ouchi N, Takahashi M, Igura T, Inui Y, Kihara S, Nakamura T, Yamashita S, Miyagawa J, Funahashi T, Matsuzawa Y 2000 An adipocyte-derived plasma protein, adiponectin, adheres to injured vascular walls. Horm Metab Res 32:47-50 197. Yamauchi T, Kamon J, Waki H, Imai Y, Shimozawa N, Hioki K, Uchida S, Ito Y, Takakuwa K, Matsui J, Takata M, Eto K, Terauchi Y, Komeda K, Tsunoda M, Murakami K, Ohnishi Y, Naitoh T, Yamamura K, Ueyama Y, Froguel P, Kimura S, Nagai R,

100 Kadowaki T 2003 Globular adiponectin protected ob/ob mice from diabetes and ApoE- deficient mice from atherosclerosis. Journal of Biological Chemistry 278:2461-2468 198. Ouchi N, Kihara S, Arita Y, Maeda K, Kuriyama H, Okamoto Y, Hotta K, Nishida M, Takahashi M, Nakamura T, Yamashita S, Funahashi T, Matsuzawa Y 1999 Novel modulator for endothelial adhesion molecules: adipocyte-derived plasma protein adiponectin. Circulation 100:2473-2476 199. Fantuzzi G 2005 Adipose tissue, adipokines, and inflammation. J Allergy Clin Immunol 115:911-919 200. Hug C, Lodish HF 2005 The role of the adipocyte hormone adiponectin in cardiovascular disease. Curr Opin Pharmacol 5:129-134 201. Degawa-Yamauchi M, Moss KA, Bovenkerk JE, Shankar SS, Morrison CL, Lelliott CJ, Vidal-Puig A, Jones R, Considine RV 2005 Regulation of adiponectin expression in human adipocytes: effects of adiposity, glucocorticoids, and tumor necrosis factor alpha. Obes Res 13:662-669 202. Kersten S, Mandard S, Tan NS, Escher P, Metzger D, Chambon P, Gonzalez FJ, Desvergne B, Wahli W 2000 Characterization of the fasting-induced adipose factor FIAF, a novel peroxisome proliferator-activated receptor target gene. Journal of Biological Chemistry 275:28488-28493 203. Kim I, Kim HG, Kim H, Kim HH, Park SK, Uhm CS, Lee ZH, Koh GY 2000 Hepatic expression, synthesis and secretion of a novel fibrinogen/angiopoietin-related protein that prevents endothelial-cell apoptosis. Biochem J 346 Pt 3:603-610 204. Yoon JC, Chickering TW, Rosen ED, Dussault B, Qin Y, Soukas A, Friedman JM, Holmes WE, Spiegelman BM 2000 Peroxisome proliferator-activated receptor gamma target gene encoding a novel angiopoietin-related protein associated with adipose differentiation. Mol Cell Biol 20:5343-5349 205. Ge H, Yang G, Huang L, Motola DL, Pourbahrami T, Li C 2004 Oligomerization and regulated proteolytic processing of angiopoietin-like protein 4. Journal of Biological Chemistry 279:2038-2045 206. Mandard S, Zandbergen F, Tan NS, Escher P, Patsouris D, Koenig W, Kleemann R, Bakker A, Veenman F, Wahli W, Muller M, Kersten S 2004 The direct peroxisome proliferator-activated receptor target fasting-induced adipose factor (FIAF/PGAR/ANGPTL4) is present in blood plasma as a truncated protein that is increased by fenofibrate treatment. Journal of Biological Chemistry 279:34411-34420 207. Yoshida K, Ono M, Koishi R, Furukawa H 2004 Characterization of the 5' regulatory region of the mouse angiopoietin-like protein 4. Vet Res Commun 28:299-305 208. Belanger AJ, Lu H, Date T, Liu LX, Vincent KA, Akita GY, Cheng SH, Gregory RJ, Jiang C 2002 Hypoxia up-regulates expression of peroxisome proliferator-activated receptor gamma angiopoietin-related gene (PGAR) in cardiomyocytes: role of hypoxia inducible factor 1alpha. J Mol Cell Cardiol 34:765-774 209. Ge H, Cha JY, Gopal H, Harp C, Yu X, Repa JJ, Li C 2005 Differential regulation and properties of angiopoietin-like proteins 3 and 4. J Lipid Res 46:1484-1490 210. Yamada T, Ozaki N, Kato Y, Miura Y, Oiso Y 2006 Insulin downregulates angiopoietin- like protein 4 mRNA in 3T3-L1 adipocytes. Biochem Biophys Res Commun 347:1138- 1144

101 211. Yoshida K, Shimizugawa T, Ono M, Furukawa H 2002 Angiopoietin-like protein 4 is a potent hyperlipidemia-inducing factor in mice and inhibitor of lipoprotein lipase. J Lipid Res 43:1770-1772 212. Ge H, Yang G, Yu X, Pourbahrami T, Li C 2004 Oligomerization state-dependent hyperlipidemic effect of angiopoietin-like protein 4. J Lipid Res 45:2071-2079 213. Koster A, Chao YB, Mosior M, Ford A, Gonzalez-DeWhitt PA, Hale JE, Li D, Qiu Y, Fraser CC, Yang DD, Heuer JG, Jaskunas SR, Eacho P 2005 Transgenic angiopoietin- like (angptl)4 overexpression and targeted disruption of angptl4 and angptl3: regulation of triglyceride metabolism. Endocrinology 146:4943-4950 214. Kersten S 2005 Regulation of lipid metabolism via angiopoietin-like proteins. Biochemal Society Transections 33:1059-1062 215. Mandard S, Zandbergen F, van Straten E, Wahli W, Kuipers F, Muller M, Kersten S 2006 The fasting-induced adipose factor/angiopoietin-like protein 4 is physically associated with lipoproteins and governs plasma lipid levels and adiposity. Journal of Biological Chemistry 281:934-944 216. Le Jan S, Amy C, Cazes A, Monnot C, Lamande N, Favier J, Philippe J, Sibony M, Gasc JM, Corvol P, Germain S 2003 Angiopoietin-like 4 is a proangiogenic factor produced during ischemia and in conventional renal cell carcinoma. Am J Pathol 162:1521-1528 217. Zhu H, Li J, Qin W, Yang Y, He X, Wan D, Gu J 2002 [Cloning of a novel gene, ANGPTL4 and the functional study in angiogenesis]. Zhonghua Yi Xue Za Zhi 82:94-99 218. Hermann LM, Pinkerton M, Jennings K, Yang L, Grom A, Sowders D, Kersten S, Witte DP, Hirsch R, Thornton S 2005 Angiopoietin-like-4 is a potential angiogenic mediator in arthritis. Clin Immunol 115:93-101 219. GOLOSOW N, GROBSTEIN C 1962 Epitheliomesenchymal interaction in pancreatic morphogenesis. Dev Biol 4:242-255 220. Bottcher G, Sjoberg J, Ekman R, Hakanson R, Sundler F 1993 Peptide YY in the mammalian pancreas: immunocytochemical localization and immunochemical characterization. Regul Pept 43:115-130 221. Rouiller DG, Cirulli V, Halban PA 1990 Differences in aggregation properties and levels of the neural cell adhesion molecule (NCAM) between islet cell types. Exp Cell Res 191:305-312 222. Slack JM 1995 Developmental biology of the pancreas. Development 121:1569-1580 223. Banting FG, Best CH 1990 Pancreatic extracts. 1922. J Lab Clin Med 115:254-272 224. Munjaal RP, Saunders GF 1979 Isolation and characterization of preproinsulin mRNA from fetal bovine pancreatic islets. Mol Cell Endocrinol 15:51-60 225. Giddings SJ, Carnaghi LR 1988 The two nonallelic rat insulin mRNAs and pre-mRNAs are regulated coordinately in vivo. Journal of Biological Chemistry 263:3845-3849 226. Steiner DF, Chan SJ, Welsh JM, Kwok SC 1985 Structure and evolution of the insulin gene. Annu Rev Genet 19:463-484 227. Melloul D, Marshak S, Cerasi E 2002 Regulation of insulin gene transcription. Diabetologia 45:309-326 228. Mosley AL, Corbett JA, Ozcan S 2004 Glucose regulation of insulin gene expression requires the recruitment of p300 by the beta-cell-specific transcription factor Pdx-1. Molecular Endocrinology 18:2279-2290

102 229. Kelpe CL, Moore PC, Parazzoli SD, Wicksteed B, Rhodes CJ, Poitout V 2003 Palmitate inhibition of insulin gene expression is mediated at the transcriptional level via ceramide synthesis. Journal of Biological Chemistry 278:30015-30021 230. Laubner K, Kieffer TJ, Lam NT, Niu X, Jakob F, Seufert J 2005 Inhibition of preproinsulin gene expression by leptin induction of suppressor of cytokine signaling 3 in pancreatic beta-cells. Diabetes 54:3410-3417 231. Zelent D, Najafi H, Odili S, Buettger C, Weik-Collins H, Li C, Doliba N, Grimsby J, Matschinsky FM 2005 Glucokinase and glucose homeostasis: proven concepts and new ideas. Biochemal Society Transections 33:306-310 232. Matschinsky FM 1996 Banting Lecture 1995. A lesson in metabolic regulation inspired by the glucokinase glucose sensor paradigm. Diabetes 45:223-241 233. Rorsman P 1997 The pancreatic beta-cell as a fuel sensor: an electrophysiologist's viewpoint. Diabetologia 40:487-495 234. Lang J 1999 Molecular mechanisms and regulation of insulin exocytosis as a paradigm of endocrine secretion. Eur J Biochem 259:3-17 235. Henquin JC 2000 Triggering and amplifying pathways of regulation of insulin secretion by glucose. Diabetes 49:1751-1760 236. Matschinsky FM, Magnuson MA, Zelent D, Jetton TL, Doliba N, Han Y, Taub R, Grimsby J 2006 The network of glucokinase-expressing cells in glucose homeostasis and the potential of glucokinase activators for diabetes therapy. Diabetes 55:1-12 237. Niwa T, Matsukawa Y, Senda T, Nimura Y, Hidaka H, Niki I 1998 Acetylcholine activates intracellular movement of insulin granules in pancreatic beta-cells via inositol trisphosphate-dependent [correction of triphosphate-dependent] mobilization of intracellular Ca2+. Diabetes 47:1699-1706 238. Baile CA, Glick Z, Mayer J 1969 Effects of secretin and cholecystokinin-pancreozymin on pancreatic juice and insulin secretion of goats. J Dairy Sci 52:513-517 239. Turner DS, Etheridge L, Jones J, Marks V, Meldrum B, Bloom SR, Brown JC 1974 The effect of the intestinal polypeptides, IRP and GIP, on insulin release and glucose tolerance in the baboon. Clin Endocrinol (Oxf) 3:489-493 240. Ahren B, Havel PJ 1999 Leptin inhibits insulin secretion induced by cellular cAMP in a pancreatic B cell line (INS-1 cells). Am J Physiol 277:R959-R966 241. Ishida K, Murakami T, Mizuno A, Iida M, Kuwajima M, Shima K 1997 Leptin suppresses basal insulin secretion from rat pancreatic islets. Regul Pept 70:179-182 242. Fehmann HC, Peiser C, Bode HP, Stamm M, Staats P, Hedetoft C, Lang RE, Goke B 1997 Leptin: a potent inhibitor of insulin secretion. Peptides 18:1267-1273 243. Persson-Sjogren S, Elmi A, Lindstrom P 2004 Effects of leptin, acetylcholine and vasoactive intestinal polypeptide on insulin secretion in isolated ob/ob mouse pancreatic islets. Acta Diabetol 41:104-112 244. Winzell MS, Nogueiras R, Dieguez C, Ahren B 2004 Dual action of adiponectin on insulin secretion in insulin-resistant mice. Biochem Biophys Res Commun 321:154-160 245. Bouwens L, Rooman I 2005 Regulation of pancreatic beta-cell mass. Physiol Rev 85:1255-1270 246. Street CN, Lakey JR, Shapiro AM, Imes S, Rajotte RV, Ryan EA, Lyon JG, Kin T, Avila J, Tsujimura T, Korbutt GS 2004 Islet graft assessment in the Edmonton Protocol: implications for predicting long-term clinical outcome. Diabetes 53:3107-3114

103 247. Brelje TC, Scharp DW, Lacy PE, Ogren L, Talamantes F, Robertson M, Friesen HG, Sorenson RL 1993 Effect of homologous placental lactogens, prolactins, and growth hormones on islet B-cell division and insulin secretion in rat, mouse, and human islets: implication for placental lactogen regulation of islet function during pregnancy. Endocrinology 132:879-887 248. Sorenson RL, Brelje TC 1997 Adaptation of islets of Langerhans to pregnancy: beta-cell growth, enhanced insulin secretion and the role of lactogenic hormones. Horm Metab Res 29:301-307 249. Blondeau B, Garofano A, Czernichow P, Breant B 1999 Age-dependent inability of the endocrine pancreas to adapt to pregnancy: a long-term consequence of perinatal malnutrition in the rat. Endocrinology 140:4208-4213 250. Larner J 1973 Purification and characterization of glycogen synthase. Birth Defects Orig Artic Ser 9:149-159 251. Pitcher J, Smythe C, Campbell DG, Cohen P 1987 Identification of the 38-kDa subunit of rabbit skeletal muscle glycogen synthase as glycogenin. Eur J Biochem 169:497-502 252. Pugazhenthi S, Khandelwal RL 1995 Regulation of glycogen synthase activation in isolated hepatocytes. Mol Cell Biochem 149-150:95-101 253. Brady MJ, Saltiel AR 2001 The role of protein phosphatase-1 in insulin action. Recent Prog Horm Res 56:157-173 254. WEBER G 1961 Kidney enzymes of gluconeogenesis, glycogenesis, glycolysis and direct oxidation. Proceedings of the Society for experimental Biology and Medicine 108:631-634 255. Katz J, Tayek JA 1999 Recycling of glucose and determination of the Cori Cycle and gluconeogenesis. Am J Physiol 277:E401-E407 256. NIKKILAE EA, OJALA K 1964 GLUCONEOGENESIS FROM GLYCEROL IN FASTING RATS. Life Sciences 3:243-249 257. Mallet LE, Exton JH, Park CR 1969 Control of gluconeogenesis from amino acids in the perfused rat liver. Journal of Biological Chemistry 244:5713-5723 258. Zammit VA 1995 Insulin and the partitioning of hepatic fatty acid metabolism. Biochemal Society Transections 23:506-511 259. Davidson NO, Shelness GS 2000 APOLIPOPROTEIN B: mRNA editing, lipoprotein assembly, and presecretory degradation. Annu Rev Nutr 20:169-193 260. Olofsson SO, Stillemark-Billton P, Asp L 2000 Intracellular assembly of VLDL: two major steps in separate cell compartments. Trends Cardiovasc Med 10:338-345 261. Boren J, White A, Wettesten M, Scott J, Graham L, Olofsson SO 1991 The molecular mechanism for the assembly and secretion of ApoB-100-containing lipoproteins. Prog Lipid Res 30:205-218 262. Havel RJ, Fielding CJ, Olivecrona T, Shore VG, Fielding PE, Egelrud T 1973 Cofactor activity of protein components of human very low density lipoproteins in the hydrolysis of triglycerides by lipoproteins lipase from different sources. Biochemistry 12:1828-1833 263. Kalra SP, Dube MG, Pu S, Xu B, Horvath TL, Kalra PS 1999 Interacting appetite- regulating pathways in the hypothalamic regulation of body weight. Endocr Rev 20:68- 100 264. Cone RD, Cowley MA, Butler AA, Fan W, Marks DL, Low MJ 2001 The arcuate nucleus as a conduit for diverse signals relevant to energy homeostasis. Int J Obes Relat Metab Disord 25 Suppl 5:S63-S67

104 265. Seeley RJ, Drazen DL, Clegg DJ 2004 The critical role of the melanocortin system in the control of energy balance. Annu Rev Nutr 24:133-149 266. Jacobowitz DM, O'Donohue TL 1978 alpha-Melanocyte stimulating hormone: immunohistochemical identification and mapping in neurons of rat brain. Proc Natl Acad Sci U S A 75:6300-6304 267. Joseph SA, Michael GJ 1988 Efferent ACTH-IR opiocortin projections from nucleus tractus solitarius: a hypothalamic deafferentation study. Peptides 9:193-201 268. Cone RD, Lu D, Koppula S, Vage DI, Klungland H, Boston B, Chen W, Orth DN, Pouton C, Kesterson RA 1996 The melanocortin receptors: agonists, antagonists, and the hormonal control of pigmentation. Recent Prog Horm Res 51:287-317 269. Kishi T, Aschkenasi CJ, Lee CE, Mountjoy KG, Saper CB, Elmquist JK 2003 Expression of melanocortin 4 receptor mRNA in the central nervous system of the rat. J Comp Neurol 457:213-235 270. Huszar D, Lynch CA, Fairchild-Huntress V, Dunmore JH, Fang Q, Berkemeier LR, Gu W, Kesterson RA, Boston BA, Cone RD, Smith FJ, Campfield LA, Burn P, Lee F 1997 Targeted disruption of the melanocortin-4 receptor results in obesity in mice. Cell 88:131-141 271. Benoit SC, Schwartz MW, Lachey JL, Hagan MM, Rushing PA, Blake KA, Yagaloff KA, Kurylko G, Franco L, Danhoo W, Seeley RJ 2000 A novel selective melanocortin-4 receptor agonist reduces food intake in rats and mice without producing aversive consequences. J Neurosci 20:3442-3448 272. Fan W, Boston BA, Kesterson RA, Hruby VJ, Cone RD 1997 Role of melanocortinergic neurons in feeding and the agouti obesity syndrome. Nature 385:165-168 273. Abbott CR, Rossi M, Kim M, AlAhmed SH, Taylor GM, Ghatei MA, Smith DM, Bloom SR 2000 Investigation of the melanocyte stimulating hormones on food intake. Lack Of evidence to support a role for the melanocortin-3-receptor. Brain Research 869:203-210 274. Haskell-Luevano C, Chen P, Li C, Chang K, Smith MS, Cameron JL, Cone RD 1999 Characterization of the neuroanatomical distribution of agouti-related protein immunoreactivity in the rhesus monkey and the rat. Endocrinology 140:1408-1415 275. Ollmann MM, Wilson BD, Yang YK, Kerns JA, Chen Y, Gantz I, Barsh GS 1997 Antagonism of central melanocortin receptors in vitro and in vivo by agouti-related protein. Science 278:135-138 276. Bagnol D, Lu XY, Kaelin CB, Day HE, Ollmann M, Gantz I, Akil H, Barsh GS, Watson SJ 1999 Anatomy of an endogenous antagonist: relationship between Agouti-related protein and proopiomelanocortin in brain. J Neurosci 19:RC26 277. Graham M, Shutter JR, Sarmiento U, Sarosi I, Stark KL 1997 Overexpression of Agrt leads to obesity in transgenic mice. Nat Genet 17:273-274 278. Small CJ, Liu YL, Stanley SA, Connoley IP, Kennedy A, Stock MJ, Bloom SR 2003 Chronic CNS administration of Agouti-related protein (Agrp) reduces energy expenditure. Int J Obes Relat Metab Disord 27:530-533 279. Goto K, Inui A, Takimoto Y, Yuzuriha H, Asakawa A, Kawamura Y, Tsuji H, Takahara Y, Takeyama C, Katsuura G, Kasuga M 2003 Acute intracerebroventricular administration of either carboxyl-terminal or amino-terminal fragments of agouti-related peptide produces a long-term decrease in energy expenditure in rats. Int J Mol Med 12:379-383

105 280. Makimura H, Mizuno TM, Mastaitis JW, Agami R, Mobbs CV 2002 Reducing hypothalamic AGRP by RNA interference increases metabolic rate and decreases body weight without influencing food intake. BMC Neurosci 3:18 281. Xiao E, Xia-Zhang L, Vulliemoz NR, Ferin M, Wardlaw SL 2003 Agouti-related protein stimulates the hypothalamic-pituitary-adrenal (HPA) axis and enhances the HPA response to interleukin-1 in the primate. Endocrinology 144:1736-1741 282. Stanley BG, Leibowitz SF 1985 Neuropeptide Y injected in the paraventricular hypothalamus: a powerful stimulant of feeding behavior. Proc Natl Acad Sci U S A 82:3940-3943 283. Erickson JC, Hollopeter G, Palmiter RD 1996 Attenuation of the obesity syndrome of ob/ob mice by the loss of neuropeptide Y. Science 274:1704-1707 284. Qian S, Chen H, Weingarth D, Trumbauer ME, Novi DE, Guan X, Yu H, Shen Z, Feng Y, Frazier E, Chen A, Camacho RE, Shearman LP, Gopal-Truter S, MacNeil DJ, Van Der Ploeg LH, Marsh DJ 2002 Neither agouti-related protein nor neuropeptide Y is critically required for the regulation of energy homeostasis in mice. Mol Cell Biol 22:5027-5035 285. Horvath TL, Diano S, Tschop M 2004 Brain circuits regulating energy homeostasis. Neuroscientist 10:235-246 286. Woods SC, Benoit SC, Clegg DJ, Seeley RJ 2004 Clinical endocrinology and metabolism. Regulation of energy homeostasis by peripheral signals. Best Pract Res Clin Endocrinol Metab 18:497-515 287. Tschop M, Smiley DL, Heiman ML 2000 Ghrelin induces adiposity in rodents. Nature 407:908-913 288. Obici S, Feng Z, Morgan K, Stein D, Karkanias G, Rossetti L 2002 Central administration of oleic acid inhibits glucose production and food intake. Diabetes 51:271- 275 289. Oomura Y, Ono T, Ooyama H, Wayner MJ 1969 Glucose and osmosensitive neurones of the rat hypothalamus. Nature 222:282-284 290. Mayer J 1955 Regulation of energy intake and the body weight: the glucostatic theory and the lipostatic hypothesis. Ann N Y Acad Sci 63:15-43 291. Grill HJ, Kaplan JM 2002 The neuroanatomical axis for control of energy balance. Front Neuroendocrinol 23:2-40 292. Schwartz MW, Woods SC, Porte D, Jr., Seeley RJ, Baskin DG 2000 Central nervous system control of food intake. Nature 404:661-671 293. Woods SC 2004 Gastrointestinal satiety signals I. An overview of gastrointestinal signals that influence food intake. Am J Physiol Gastrointest Liver Physiol 286:G7-13 294. Matson CA, Wiater MF, Kuijper JL, Weigle DS 1997 Synergy between leptin and cholecystokinin (CCK) to control daily caloric intake. Peptides 18:1275-1278 295. Riedy CA, Chavez M, Figlewicz DP, Woods SC 1995 Central insulin enhances sensitivity to cholecystokinin. Physiol Behav 58:755-760 296. Stubbs RJ 1995 Macronutrient effects on appetite. Int J Obes Relat Metab Disord 19 Suppl 5:S11-S19 297. Holmes LJ, Storlien LH, Smythe GA 1989 Hypothalamic monoamines associated with the cephalic phase insulin response. Am J Physiol 256:E236-E241

106 298. Coppack SW, Fisher RM, Gibbons GF, Humphreys SM, McDonough MJ, Potts JL, Frayn KN 1990 Postprandial substrate deposition in human forearm and adipose tissues in vivo. Clin Sci (Lond) 79:339-348 299. Yu YH, Ginsberg HN 2005 Adipocyte signaling and lipid homeostasis: sequelae of insulin-resistant adipose tissue. Circ Res 96:1042-1052 300. Evans K, Burdge GC, Wootton SA, Clark ML, Frayn KN 2002 Regulation of dietary fatty acid entrapment in subcutaneous adipose tissue and skeletal muscle. Diabetes 51:2684-2690 301. Schutz Y 2004 Dietary fat, lipogenesis and energy balance. Physiol Behav 83:557-564 302. Havel PJ, Townsend R, Chaump L, Teff K 1999 High-fat meals reduce 24-h circulating leptin concentrations in women. Diabetes 48:334-341 303. Kaiyala KJ, Prigeon RL, Kahn SE, Woods SC, Schwartz MW 2000 Obesity induced by a high-fat diet is associated with reduced brain insulin transport in dogs. Diabetes 49:1525- 1533 304. Kahn BB, Cushman SW, Flier JS 1989 Regulation of glucose transporter-specific mRNA levels in rat adipose cells with fasting and refeeding. Implications for in vivo control of glucose transporter number. J Clin Invest 83:199-204 305. Moran O, Phillip M 2003 Leptin: obesity, diabetes and other peripheral effects--a review. Pediatr Diabetes 4:101-109 306. Caro JF, Kolaczynski JW, Nyce MR, Ohannesian JP, Opentanova I, Goldman WH, Lynn RB, Zhang PL, Sinha MK, Considine RV 1996 Decreased cerebrospinal-fluid/serum leptin ratio in obesity: a possible mechanism for leptin resistance. Lancet 348:159-161 307. Edgerton DS, Jacobson PB, Opgenorth TJ, Zinker B, Beno D, von Geldern T, Ohman L, Scott M, Neal D, Cherrington AD 2006 Selective antagonism of the hepatic glucocorticoid receptor reduces hepatic glucose production. Metabolism 55:1255-1262 308. Nonogaki K 2000 New insights into sympathetic regulation of glucose and fat metabolism. Diabetologia 43:533-549 309. Tempel DL, McEwen BS, Leibowitz SF 1992 Effects of adrenal steroid agonists on food intake and macronutrient selection. Physiol Behav 52:1161-1166 310. Strack AM, Sebastian RJ, Schwartz MW, Dallman MF 1995 Glucocorticoids and insulin: reciprocal signals for energy balance. Am J Physiol 268:R142-R149 311. York DA 1996 Lessons from animal models of obesity. Endocrinol Metab Clin North Am 25:781-800 312. Muller MJ, Seitz HJ 1984 Thyroid hormone action on intermediary metabolism. Part II: Lipid metabolism in hypo- and hyperthyroidism. Klin Wochenschr 62:49-55 313. Havel PJ 2001 Peripheral signals conveying metabolic information to the brain: short- term and long-term regulation of food intake and energy homeostasis. Exp Biol Med (Maywood ) 226:963-977 314. Luo L, MacLean DB 2003 Effects of thyroid hormone on food intake, hypothalamic Na/K ATPase activity and ATP content. Brain Research 973:233-239 315. Plata-Salaman CR 1995 Cytokines and feeding suppression: an integrative view from neurologic to molecular levels. Nutrition 11:674-677 316. Catt KJ, Moffat B, Niall HD 1967 Human growth hormone and placental lactogen: structural similarity. Science 157:321

107 317. Niall HD, Hogan ML, Sauer R, Rosenblum IY, Greenwood FC 1971 Sequences of pituitary and placental lactogenic and growth hormones: evolution from a primordial peptide by gene reduplication. Proc Natl Acad Sci U S A 68:866-870 318. Ono M, Takayama Y, Rand-Weaver M, Sakata S, Yasunaga T, Noso T, Kawauchi H 1990 cDNA cloning of somatolactin, a pituitary protein related to growth hormone and prolactin. Proc Natl Acad Sci U S A 87:4330-4334 319. Gubbins EJ, Maurer RA, Lagrimini M, Erwin CR, Donelson JE 1980 Structure of the rat prolactin gene. Journal of Biological Chemistry 255:8655-8662 320. Gourdji D, Laverriere JN 1994 The rat prolactin gene: a target for tissue-specific and hormone-dependent transcription factors. Mol Cell Endocrinol 100:133-142 321. Nelson C, Albert VR, Elsholtz HP, Lu LI, Rosenfeld MG 1988 Activation of cell-specific expression of rat growth hormone and prolactin genes by a common transcription factor. Science 239:1400-1405 322. Owerbach D, Rutter WJ, Cooke NE, Martial JA, Shows TB 1981 The prolactin gene is located on chromosome 6 in humans. Science 212:815-816 323. Gellersen B, DiMattia GE, Friesen HG, Bohnet HG 1989 Prolactin (PRL) mRNA from human decidua differs from pituitary PRL mRNA but resembles the IM-9-P3 lymphoblast PRL transcript. Mol Cell Endocrinol 64:127-130 324. Berwaer M, Martial JA, Davis JR 1994 Characterization of an up-stream promoter directing extrapituitary expression of the human prolactin gene. Molecular Endocrinology 8:635-642 325. Ben Jonathan N, Mershon JL, Allen DL, Steinmetz RW 1996 Extrapituitary prolactin: distribution, regulation, functions, and clinical aspects. Endocr Rev 17:639-669 326. Nicoll CS, Mayer GL, Russell SM 1986 Structural features of prolactins and growth hormones that can be related to their biological properties. Endocr Rev 7:169-203 327. Sinha YN 1995 Structural variants of prolactin: occurrence and physiological significance. Endocr Rev 16:354-369 328. Keeler C, Dannies PS, Hodsdon ME 2003 The tertiary structure and backbone dynamics of human prolactin. J Mol Biol 328:1105-1121 329. Markoff E, Sigel MB, Lacour N, Seavey BK, Friesen HG, Lewis UJ 1988 Glycosylation selectively alters the biological activity of prolactin. Endocrinology 123:1303-1306 330. Ho TW, Leong FS, Olaso CH, Walker AM 1993 Secretion of specific nonphosphorylated and phosphorylated rat prolactin isoforms at different stages of the estrous cycle. Neuroendocrinology 58:160-165 331. Kim BG, Brooks CL 1993 Isolation and characterization of phosphorylated bovine prolactin. Biochem J 296 ( Pt 1):41-47 332. Oetting WS, Tuazon PT, Traugh JA, Walker AM 1986 Phosphorylation of prolactin. Journal of Biological Chemistry 261:1649-1652 333. Wicks JR, Brooks CL 1995 Biological activity of phosphorylated and dephosphorylated bovine prolactin. Mol Cell Endocrinol 112:223-229 334. Ueda E, Ozerdem U, Chen YH, Yao M, Huang KT, Sun H, Martins-Green M, Bartolini P, Walker AM 2006 A molecular mimic demonstrates that phosphorylated human prolactin is a potent anti-angiogenic hormone. Endocr Relat Cancer 13:95-111 335. Clapp C, Sears PS, Russell DH, Richards J, Levay-Young BK, Nicoll CS 1988 Biological and immunological characterization of cleaved and 16K forms of rat prolactin. Endocrinology 122:2892-2898

108 336. Clapp C, Martial JA, Guzman RC, Rentier-Delure F, Weiner RI 1993 The 16-kilodalton N-terminal fragment of human prolactin is a potent inhibitor of angiogenesis. Endocrinology 133:1292-1299 337. Corbacho AM, Martinez DLE, Clapp C 2002 Roles of prolactin and related members of the prolactin/growth hormone/placental lactogen family in angiogenesis. Journal of Endocrinology 173:219-238 338. Herlant M 1964 The cells of the adenohypophysis and their functional significance. Int Rev Cytol 17:299-382 339. Baker BL, Gross DS 1978 Cytology and distribution of secretory cell types in the mouse hypophysis as demonstrated with immunocytochemistry. Am J Anat 153:193-215 340. Halmi NS, Parsons JA, Erlandsen SL, Duello T 1975 Prolactin and growth hormone cells in the human hypophysis: a study with immunoenzyme histochemistry and differential staining. Cell Tissue Res 158:497-507 341. Scully KM, Rosenfeld MG 2002 Pituitary development: regulatory codes in mammalian organogenesis. Science 295:2231-2235 342. Li S, Crenshaw EB, III, Rawson EJ, Simmons DM, Swanson LW, Rosenfeld MG 1990 Dwarf locus mutants lacking three pituitary cell types result from mutations in the POU- domain gene pit-1. Nature 347:528-533 343. Lin SC, Lin CR, Gukovsky I, Lusis AJ, Sawchenko PE, Rosenfeld MG 1993 Molecular basis of the little mouse phenotype and implications for cell type-specific growth. Nature 364:208-213 344. Chen HT 1987 Postnatal development of pituitary lactotropes in the rat measured by reverse hemolytic plaque assay. Endocrinology 120:247-253 345. De Paul AL, Pons P, Aoki A, Torres AI 1997 Heterogeneity of pituitary lactotrophs: immunocytochemical identification of functional subtypes. Acta Histochem 99:277-289 346. Nakane PK 1970 Classifications of anterior pituitary cell types with immunoenzyme histochemistry. J Histochem Cytochem 18:9-20 347. Boockfor FR, Frawley LS 1987 Functional variations among prolactin cells from different pituitary regions. Endocrinology 120:874-879 348. Lledo PM, Guerineau N, Mollard P, Vincent JD, Israel JM 1991 Physiological characterization of two functional states in subpopulations of prolactin cells from lactating rats. J Physiol 437:477-494 349. Lewis DL, Goodman MB, St John PA, Barker JL 1988 Calcium currents and fura-2 signals in fluorescence-activated cell sorted lactotrophs and somatotrophs of rat anterior pituitary. Endocrinology 123:611-621 350. Frawley LS, Boockfor FR, Hoeffler JP 1985 Identification by plaque assays of a pituitary cell type that secretes both growth hormone and prolactin. Endocrinology 116:734-737 351. Frawley LS, Boockfor FR 1991 Mammosomatotropes: presence and functions in normal and neoplastic pituitary tissue. Endocr Rev 12:337-355 352. Boockfor FR, Hoeffler JP, Frawley LS 1986 Estradiol induces a shift in cultured cells that release prolactin or growth hormone. Am J Physiol 250:E103-E105 353. Ling C, Svensson L, Oden B, Weijdegard B, Eden B, Eden S, Billig H 2003 Identification of functional prolactin (PRL) receptor gene expression: PRL inhibits lipoprotein lipase activity in human white adipose tissue. J Clin Endocrinol Metab 88:1804-1808

109 354. Hugo ER, Brandebourg TD, Comstock CE, Gersin KS, Sussman JJ, Ben Jonathan N 2006 LS14: a novel human adipocyte cell line that produces prolactin. Endocrinology 147:306-313 355. Handwerger S, Freemark M 2000 The roles of placental growth hormone and placental lactogen in the regulation of human fetal growth and development. J Pediatr Endocrinol Metab 13:343-356 356. Tashjian AH, Jr., Yasumura Y, Levine L, Sato GH, Parker ML 1968 Establishment of clonal strains of rat pituitary tumor cells that secrete growth hormone. Endocrinology 82:342-352 357. Judd AM, Login IS, Kovacs K, Ross PC, Spangelo BL, Jarvis WD, MacLeod RM 1988 Characterization of the MMQ cell, a prolactin-secreting clonal cell line that is responsive to dopamine. Endocrinology 123:2341-2350 358. Pastorcic M, De A, Boyadjieva N, Vale W, Sarkar DK 1995 Reduction in the expression and action of transforming growth factor beta 1 on lactotropes during estrogen-induced tumorigenesis in the anterior pituitary. Cancer Res 55:4892-4898 359. Kamberi IA, Mical RS, Porter JC 1970 Prolactin-inhibiting activity in hypophysial stalk blood and elevation by dopamine. Experientia 26:1150-1151 360. Bishop W, Krulich L, Fawcett CP, McCann SM 1971 The effect of median eminence (ME) lesions on plasma levels of FSH, LH, and prolactin in the rat. Proceedings of the Society for experimental Biology and Medicine 136:925-927 361. Ben Jonathan N 1985 Dopamine: a prolactin-inhibiting hormone. Endocr Rev 6:564-589 362. Ben Jonathan N, Oliver C, Weiner HJ, Mical RS, Porter JC 1977 Dopamine in hypophysial portal plasma of the rat during the estrous cycle and throughout pregnancy. Endocrinology 100:452-458 363. Arbogast LA, Voogt JL 1991 Hyperprolactinemia increases and hypoprolactinemia decreases tyrosine hydroxylase messenger ribonucleic acid levels in the arcuate nuclei, but not the substantia nigra or zona incerta. Endocrinology 128:997-1005 364. Ben Jonathan N, Peters LL 1982 Posterior pituitary lobectomy: differential elevation of plasma prolactin and luteinizing hormone in estrous and lactating rats. Endocrinology 110:1861-1865 365. Peters LL, Hoefer MT, Ben Jonathan N 1981 The posterior pituitary: regulation of anterior pituitary prolactin secretion. Science 213:659-661 366. Minami S, Sarkar DK 1997 Transforming growth factor-beta 1 inhibits prolactin secretion and lactotropic cell proliferation in the pituitary of oestrogen-treated Fischer 344 rats. Neurochem Int 30:499-506 367. Shah GV, Wang W, Grosvenor CE, Crowley WR 1990 Calcitonin inhibits basal and thyrotropin-releasing hormone-induced release of prolactin from anterior pituitary cells: evidence for a selective action exerted proximal to secretagogue-induced increases in cytosolic Ca2+. Endocrinology 127:621-628 368. Tohei A, VandeGarde B, Arbogast LA, Voogt JL 2000 Calcitonin inhibition of prolactin secretion in lactating rats: mechanism of action. Neuroendocrinology 71:327-332 369. Nakayama Y, Hattori N, Otani H, Inagaki C 2006 Gamma-aminobutyric acid (GABA)-C receptor stimulation increases prolactin (PRL) secretion in cultured rat anterior pituitary cells. Biochem Pharmacol 71:1705-1710

110 370. Lledo PM, Legendre P, Israel JM, Vincent JD 1990 Dopamine inhibits two characterized voltage-dependent calcium currents in identified rat lactotroph cells. Endocrinology 127:990-1001 371. Ben Jonathan N, Hnasko R 2001 Dopamine as a prolactin (PRL) inhibitor. Endocr Rev 22:724-763 372. Arita J, Hashi A, Hoshi K, Mazawa S, Suzuki S 1998 D2 dopamine-receptor-mediated inhibition of proliferation of rat lactotropes in culture is accompanied by changes in cell shape. Neuroendocrinology 68:163-171 373. Blake CA 1974 Stimulation of pituitary prolactin and TSH release in lactating and proestrous rats. Endocrinology 94:503-508 374. Tashjian AH, Jr., Barowsky NJ, Jensen DK 1971 Thyrotropin releasing hormone: direct evidence for stimulation of prolactin production by pituitary cells in culture. Biochem Biophys Res Commun 43:516-523 375. Deis RP, Alonso N 1973 Prolactin release induced by synthetic thyrotrophin releasing factor in female rats. Journal of Endocrinology 58:673-674 376. Fomina AF, Levitan ES 1995 Three phases of TRH-induced facilitation of exocytosis by single lactotrophs. J Neurosci 15:4982-4991 377. Ruberg M, Rotsztejn WH, Arancibia S, Besson J, Enjalbert A 1978 Stimulation of prolactin release by vasoactive intestinal peptide (VIP). Eur J Pharmacol 51:319-320 378. Reichlin S 1988 Neuroendocrine significance of vasoactive intestinal polypeptide. Ann N Y Acad Sci 527:431-449 379. Gibbs DM 1984 High concentrations of oxytocin in hypophysial portal plasma. Endocrinology 114:1216-1218 380. Freeman ME, Kanyicska B, Lerant A, Nagy G 2000 Prolactin: structure, function, and regulation of secretion. Physiol Rev 80:1523-1631 381. Grosvenor CE, Whitworth N 1974 Evidence for a steady rate of secretion of prolactin following suckling in the rat. J Dairy Sci 57:900-904 382. Chiocchio SR, Cannata MA, Funes JR, Tramezzani JH 1979 Involvement of adenohypophysial dopamine in the regulation of prolactin release during suckling. Endocrinology 105:544-547 383. Grosvenor CE, Mena F, Whitworth NS 1977 Metabolic clearance of rat prolactin in the lactating and non-lactating rat. Journal of Endocrinology 73:1-10 384. Bodnar I, Banky ZS, Toth BE, Nagy GM, Halasz B 2002 Brain structures mediating the suckling stimulus-induced release of prolactin. J Neuroendocrinol 14:384-396 385. Zelena D, Makara GB, Nagy GM 2003 Effect of glutamate receptor antagonists on suckling-induced prolactin release in rats. Endocrine 21:147-152 386. Neill JD, Freeman ME, Tillson SA 1971 Control of the proestrus surge of prolactin and luteinizing hormone secretion by estrogens in the rat. Endocrinology 89:1448-1453 387. Arbogast LA, Ben Jonathan N 1988 The preovulatory prolactin surge: an evaluation of the role of dopamine. Endocrinology 123:2690-2695 388. Arbogast LA, Voogt JL 2002 Progesterone induces dephosphorylation and inactivation of tyrosine hydroxylase in rat hypothalamic dopaminergic neurons. Neuroendocrinology 75:273-281 389. Bertram R, Egli M, Toporikova N, Freeman ME 2006 A mathematical model for the mating-induced prolactin rhythm of female rats. Am J Physiol Endocrinol Metab 290:E573-E582

111 390. Arbogast LA, Voogt JL 1996 The responsiveness of tuberoinfundibular dopaminergic neurons to prolactin feedback is diminished between early lactation and midlactation in the rat. Endocrinology 137:47-54 391. Epstein MT, McNeilly AS, Murray MA, Hockaday TD 1975 Plasma testosterone and prolactin in the menstrual cycle. Clin Endocrinol (Oxf) 4:531-535 392. Egyed J, Doszpod J, Gati I 1978 Serum prolactin and estrogen pattern in human gestation. Endocrinol Exp 12:109-114 393. Rigg LA, Lein A, Yen SS 1977 Pattern of increase in circulating prolactin levels during human gestation. Am J Obstet Gynecol 129:454-456 394. Golander A, Hurley T, Barrett J, Hizi A, Handwerger S 1978 Prolactin synthesis by human chorion-decidual tissue: a possible source of prolactin in the amniotic fluid. Science 202:311-313 395. Winters AJ, Colston C, MacDonald PC, Porter JC 1975 Fetal plasma prolactin levels. J Clin Endocrinol Metab 41:626-629 396. Aubert ML, Grumbach MM, Kaplan SL 1974 Heterologous radioimmunoassay for plasma human prolactin (hPRL); values in normal subjects, puberty, pregnancy and in pituitary disorders. Acta Endocrinol (Copenh) 77:460-476 397. Aono T, Shioji T, Shoda T, Kurachi K 1977 The initiation of human lactation and prolactin response to suckling. J Clin Endocrinol Metab 44:1101-1106 398. Spiegel K, Follenius M, Simon C, Saini J, Ehrhart J, Brandenberger G 1994 Prolactin secretion and sleep. Sleep 17:20-27 399. Kelly PA, Djiane J, Postel-Vinay MC, Edery M 1991 The prolactin/growth hormone receptor family. Endocr Rev 12:235-251 400. Goffin V, Kelly PA 1996 Prolactin and growth hormone receptors. Clin Endocrinol (Oxf) 45:247-255 401. Bole-Feysot C, Goffin V, Edery M, Binart N, Kelly PA 1998 Prolactin (PRL) and its receptor: actions, signal transduction pathways and phenotypes observed in PRL receptor knockout mice. Endocr Rev 19:225-268 402. Buteau H, Pezet A, Ferrag F, Perrot-Applanat M, Kelly PA, Edery M 1998 N- glycosylation of the prolactin receptor is not required for activation of gene transcription but is crucial for its cell surface targeting. Molecular Endocrinology 12:544-555 403. Lebrun JJ, Ali S, Ullrich A, Kelly PA 1995 Proline-rich sequence-mediated Jak2 association to the prolactin receptor is required but not sufficient for signal transduction. Journal of Biological Chemistry 270:10664-10670 404. Ren R, Mayer BJ, Cicchetti P, Baltimore D 1993 Identification of a ten-amino acid proline-rich SH3 binding site. Science 259:1157-1161 405. Pezet A, Buteau H, Kelly PA, Edery M 1997 The last proline of Box 1 is essential for association with JAK2 and functional activation of the prolactin receptor. Mol Cell Endocrinol 129:199-208 406. Clarke DL, Linzer DI 1993 Changes in prolactin receptor expression during pregnancy in the mouse ovary. Endocrinology 133:224-232 407. Boutin JM, Jolicoeur C, Okamura H, Gagnon J, Edery M, Shirota M, Banville D, Dusanter-Fourt I, Djiane J, Kelly PA 1988 Cloning and expression of the rat prolactin receptor, a member of the growth hormone/prolactin receptor gene family. Cell 53:69-77

112 408. Shirota M, Banville D, Ali S, Jolicoeur C, Boutin JM, Edery M, Djiane J, Kelly PA 1990 Expression of two forms of prolactin receptor in rat ovary and liver. Molecular Endocrinology 4:1136-1143 409. Boutin JM, Edery M, Shirota M, Jolicoeur C, Lesueur L, Ali S, Gould D, Djiane J, Kelly PA 1989 Identification of a cDNA encoding a long form of prolactin receptor in human hepatoma and breast cancer cells. Molecular Endocrinology 3:1455-1461 410. Hu ZZ, Meng J, Dufau ML 2001 Isolation and characterization of two novel forms of the human prolactin receptor generated by alternative splicing of a newly identified exon 11. Journal of Biological Chemistry 276:41086-41094 411. Kline JB, Roehrs H, Clevenger CV 1999 Functional characterization of the intermediate isoform of the human prolactin receptor. Journal of Biological Chemistry 274:35461- 35468 412. Amit T, Dibner C, Barkey RJ 1997 Characterization of prolactin- and growth hormone- binding proteins in milk and their diversity among species. Mol Cell Endocrinol 130:167- 180 413. Postel-Vinay MC 1996 Growth hormone- and prolactin-binding proteins: soluble forms of receptors. Horm Res 45:178-181 414. Dannies PS 2001 A serum prolactin-binding protein: implications for growth hormone. Trends Endocrinol Metab 12:427-428 415. Kline JB, Clevenger CV 2001 Identification and characterization of the prolactin-binding protein in human serum and milk. Journal of Biological Chemistry 276:24760-24766 416. Ormandy CJ, Binart N, Helloco C, Kelly PA 1998 Mouse prolactin receptor gene: genomic organization reveals alternative promoter usage and generation of isoforms via alternative 3'-exon splicing. DNA Cell Biol 17:761-770 417. Moldrup A, Ormandy C, Nagano M, Murthy K, Banville D, Tronche F, Kelly PA 1996 Differential promoter usage in prolactin receptor gene expression: hepatocyte nuclear factor 4 binds to and activates the promoter preferentially active in the liver. Molecular Endocrinology 10:661-671 418. Hu Z, Zhuang L, Dufau ML 1996 Multiple and tissue-specific promoter control of gonadal and non-gonadal prolactin receptor gene expression. Journal of Biological Chemistry 271:10242-10246 419. Hu Z, Zhuang L, Guan X, Meng J, Dufau ML 1997 Steroidogenic factor-1 is an essential transcriptional activator for gonad-specific expression of promoter I of the rat prolactin receptor gene. Journal of Biological Chemistry 272:14263-14271 420. Hu ZZ, Zhuang L, Meng J, Dufau ML 1998 Transcriptional regulation of the generic promoter III of the rat prolactin receptor gene by C/EBPbeta and Sp1. Journal of Biological Chemistry 273:26225-26235 421. Arden KC, Boutin JM, Djiane J, Kelly PA, Cavenee WK 1990 The receptors for prolactin and growth hormone are localized in the same region of human chromosome 5. Cytogenet Cell Genet 53:161-165 422. Hu ZZ, Zhuang L, Meng J, Tsai-Morris CH, Dufau ML 2002 Complex 5' genomic structure of the human prolactin receptor: multiple alternative exons 1 and promoter utilization. Endocrinology 143:2139-2142 423. Hu ZZ, Zhuang L, Meng J, Leondires M, Dufau ML 1999 The human prolactin receptor gene structure and alternative promoter utilization: the generic promoter hPIII and a novel human promoter hP(N). J Clin Endocrinol Metab 84:1153-1156

113 424. Goffin V, Martial JA, Summers NL 1995 Use of a model to understand prolactin and growth hormone specificities. Protein Eng 8:1215-1231 425. Gertler A, Petridou B, Kriwi GG, Djiane J 1993 Interaction of lactogenic hormones with purified recombinant extracellular domain of rabbit prolactin receptor expressed in insect cells. FEBS Lett 319:277-281 426. Gertler A, Grosclaude J, Strasburger CJ, Nir S, Djiane J 1996 Real-time kinetic measurements of the interactions between lactogenic hormones and prolactin-receptor extracellular domains from several species support the model of hormone-induced transient receptor dimerization. Journal of Biological Chemistry 271:24482-24491 427. Rui H, Djeu JY, Evans GA, Kelly PA, Farrar WL 1992 Prolactin receptor triggering. Evidence for rapid tyrosine kinase activation. Journal of Biological Chemistry 267:24076-24081 428. Ihle JN 1995 The Janus protein tyrosine kinase family and its role in cytokine signaling. Adv Immunol 60:1-35 429. Ihle JN 1996 STATs: signal transducers and activators of transcription. Cell 84:331-334 430. Campbell GS, Argetsinger LS, Ihle JN, Kelly PA, Rillema JA, Carter-Su C 1994 Activation of JAK2 tyrosine kinase by prolactin receptors in Nb2 cells and mouse mammary gland explants. Proc Natl Acad Sci U S A 91:5232-5236 431. Chang WP, Clevenger CV 1996 Modulation of growth factor receptor function by isoform heterodimerization. Proc Natl Acad Sci U S A 93:5947-5952 432. Horseman ND, Yu-Lee LY 1994 Transcriptional regulation by the helix bundle peptide hormones: growth hormone, prolactin, and hematopoietic cytokines. Endocr Rev 15:627- 649 433. Liu X, Robinson GW, Wagner KU, Garrett L, Wynshaw-Boris A, Hennighausen L 1997 Stat5a is mandatory for adult mammary gland development and lactogenesis. Genes and Development 11:179-186 434. Udy GB, Towers RP, Snell RG, Wilkins RJ, Park SH, Ram PA, Waxman DJ, Davey HW 1997 Requirement of STAT5b for sexual dimorphism of body growth rates and liver gene expression. Proc Natl Acad Sci U S A 94:7239-7244 435. Teglund S, McKay C, Schuetz E, van Deursen JM, Stravopodis D, Wang D, Brown M, Bodner S, Grosveld G, Ihle JN 1998 Stat5a and Stat5b proteins have essential and nonessential, or redundant, roles in cytokine responses. Cell 93:841-850 436. Buckley AR, Rao YP, Buckley DJ, Gout PW 1994 Prolactin-induced phosphorylation and nuclear translocation of MAP kinase in Nb2 lymphoma cells. Biochem Biophys Res Commun 204:1158-1164 437. Van Coppenolle F, Skryma R, Ouadid-Ahidouch H, Slomianny C, Roudbaraki M, Delcourt P, Dewailly E, Humez S, Crepin A, Gourdou I, Djiane J, Bonnal JL, Mauroy B, Prevarskaya N 2004 Prolactin stimulates cell proliferation through a long form of prolactin receptor and K+ channel activation. Biochem J 377:569-578 438. Fresno Vara JA, Carretero MV, Geronimo H, Ballmer-Hofer K, Martin-Perez J 2000 Stimulation of c-Src by prolactin is independent of Jak2. Biochem J 345 Pt 1:17-24 439. Acosta JJ, Munoz RM, Gonzalez L, Subtil-Rodriguez A, Dominguez-Caceres MA, Garcia-Martinez JM, Calcabrini A, Lazaro-Trueba I, Martin-Perez J 2003 Src mediates prolactin-dependent proliferation of T47D and MCF7 cells via the activation of focal adhesion kinase/Erk1/2 and phosphatidylinositol 3-kinase pathways. Molecular Endocrinology 17:2268-2282

114 440. Clevenger CV, Medaglia MV 1994 The protein tyrosine kinase P59fyn is associated with prolactin (PRL) receptor and is activated by PRL stimulation of T-lymphocytes. Molecular Endocrinology 8:674-681 441. Amaral ME, Cunha DA, Anhe GF, Ueno M, Carneiro EM, Velloso LA, Bordin S, Boschero AC 2004 Participation of prolactin receptors and phosphatidylinositol 3-kinase and MAP kinase pathways in the increase in pancreatic islet mass and sensitivity to glucose during pregnancy. Journal of Endocrinology 183:469-476 442. Larsen L, Ropke C 2002 Suppressors of cytokine signalling: SOCS. APMIS 110:833-844 443. Hilton DJ 1999 Negative regulators of cytokine signal transduction. Cell Mol Life Sci 55:1568-1577 444. Yoshimura A 2005 Negative regulation of cytokine signaling. Clin Rev Allergy Immunol 28:205-220 445. Tomic S, Chughtai N, Ali S 1999 SOCS-1, -2, -3: selective targets and functions downstream of the prolactin receptor. Mol Cell Endocrinol 158:45-54 446. Dif F, Saunier E, Demeneix B, Kelly PA, Edery M 2001 Cytokine-inducible SH2- containing protein suppresses PRL signaling by binding the PRL receptor. Endocrinology 142:5286-5293 447. Marine JC, Topham DJ, McKay C, Wang D, Parganas E, Stravopodis D, Yoshimura A, Ihle JN 1999 SOCS1 deficiency causes a lymphocyte-dependent perinatal lethality. Cell 98:609-616 448. Metcalf D, Greenhalgh CJ, Viney E, Willson TA, Starr R, Nicola NA, Hilton DJ, Alexander WS 2000 Gigantism in mice lacking suppressor of cytokine signalling-2. Nature 405:1069-1073 449. Marine JC, McKay C, Wang D, Topham DJ, Parganas E, Nakajima H, Pendeville H, Yasukawa H, Sasaki A, Yoshimura A, Ihle JN 1999 SOCS3 is essential in the regulation of fetal liver erythropoiesis. Cell 98:617-627 450. Ling C, Hellgren G, Gebre-Medhin M, Dillner K, Wennbo H, Carlsson B, Billig H 2000 Prolactin (PRL) receptor gene expression in mouse adipose tissue: increases during lactation and in PRL-transgenic mice. Endocrinology 141:3564-3572 451. Djiane J, Durand P, Kelly PA 1977 Evolution of prolactin receptors in rabbit mammary gland during pregnancy and lactation. Endocrinology 100:1348-1356 452. Hayden TJ, Bonney RC, Forsyth IA 1979 Ontogeny and control of prolactin receptors in the mammary gland and liver of virgin, pregnant and lactating rats. Journal of Endocrinology 80:259-269 453. Camarillo IG, Thordarson G, Moffat JG, Van Horn KM, Binart N, Kelly PA, Talamantes F 2001 Prolactin receptor expression in the epithelia and stroma of the rat mammary gland. Journal of Endocrinology 171:85-95 454. Djiane J, Durand P 1977 Prolactin-progesterone antagonism in self regulation of prolactin receptors in the mammary gland. Nature 266:641-643 455. Pi X, Grattan DR 1999 Expression of prolactin receptor mRNA is increased in the preoptic area of lactating rats. Endocrine 11:91-98 456. Bakowska JC, Morrell JI 1997 Atlas of the neurons that express mRNA for the long form of the prolactin receptor in the forebrain of the female rat. J Comp Neurol 386:161-177 457. Grattan DR, Pi XJ, Andrews ZB, Augustine RA, Kokay IC, Summerfield MR, Todd B, Bunn SJ 2001 Prolactin receptors in the brain during pregnancy and lactation: implications for behavior. Horm Behav 40:115-124

115 458. Tseng L, Zhu HH 1998 Progestin, estrogen, and insulin-like growth factor-I stimulate the prolactin receptor mRNA in human endometrial stromal cells. J Soc Gynecol Investig 5:149-155 459. Bridges RS, Hays LE 2005 Steroid-induced alterations in mRNA expression of the long form of the prolactin receptor in the medial preoptic area of female rats: Effects of exposure to a pregnancy-like regimen of progesterone and estradiol. Brain Res Mol Brain Res 140:10-16 460. Pi X, Zhang B, Li J, Voogt JL 2003 Promoter usage and estrogen regulation of prolactin receptor gene in the brain of the female rat. Neuroendocrinology 77:187-197 461. Sakai S, Banerjee MR 1979 Glucocorticoid modulation of prolactin receptors on mammary cells of lactating mice. Biochim Biophys Acta 582:79-88 462. Phillips ID, Anthony RV, Butler TG, Ross JT, McMillen IC 1997 Hepatic prolactin receptor gene expression increases in the sheep fetus before birth and after cortisol infusion. Endocrinology 138:1351-1354 463. Cassy S, Charlier M, Belair L, Guillomot M, Laud K, Djiane J 2000 Increase in prolactin receptor (PRL-R) mRNA level in the mammary gland after hormonal induction of lactation in virgin ewes. Domest Anim Endocrinol 18:41-55 464. Varas SM, Jahn GA 2005 The expression of estrogen, prolactin, and progesterone receptors in mammary gland and liver of female rats during pregnancy and early postpartum: regulation by thyroid hormones. Endocr Res 31:357-370 465. Nevalainen MT, Valve EM, Ingleton PM, Harkonen PL 1996 Expression and hormone regulation of prolactin receptors in rat dorsal and lateral prostate. Endocrinology 137:3078-3088 466. Tucker HA 2000 Hormones, mammary growth, and lactation: a 41-year perspective. J Dairy Sci 83:874-884 467. Ceriani RL 1974 Proceedings: Hormones and other factors controlling growth in the mammary gland: a review. J Invest Dermatol 63:93-108 468. Ormandy CJ, Binart N, Kelly PA 1997 Mammary gland development in prolactin receptor knockout mice. J Mammary Gland Biol Neoplasia 2:355-364 469. Horseman ND, Zhao W, Montecino-Rodriguez E, Tanaka M, Nakashima K, Engle SJ, Smith F, Markoff E, Dorshkind K 1997 Defective mammopoiesis, but normal hematopoiesis, in mice with a targeted disruption of the prolactin gene. EMBO J 16:6926-6935 470. Nandi S 1958 Endocrine control of mammarygland development and function in the C3H/ He Crgl mouse. J Natl Cancer Inst 21:1039-1063 471. AHREN K, HAMBERGER L 1962 Direct action of testosterone propionate on the rat mammary gland. Acta Endocrinol (Copenh) 40:265-276 472. Brisken C, Kaur S, Chavarria TE, Binart N, Sutherland RL, Weinberg RA, Kelly PA, Ormandy CJ 1999 Prolactin controls mammary gland development via direct and indirect mechanisms. Dev Biol 210:96-106 473. Ways J, Markoff E, Ogren L, Talamantes F 1979 Lactogenic response of mouse mammary explants from different days of pregnancy to placental lactogen and pituitary prolactin. In Vitro 15:891-894 474. Li M, Liu X, Robinson G, Bar-Peled U, Wagner KU, Young WS, Hennighausen L, Furth PA 1997 Mammary-derived signals activate programmed cell death during the first stage of mammary gland involution. Proc Natl Acad Sci U S A 94:3425-3430

116 475. Humphreys RC, Hennighausen L 1999 Signal transducer and activator of transcription 5a influences mammary epithelial cell survival and tumorigenesis. Cell Growth Differ 10:685-694 476. Capuco AV, Li M, Long E, Ren S, Hruska KS, Schorr K, Furth PA 2002 Concurrent pregnancy retards mammary involution: effects on apoptosis and proliferation of the mammary epithelium after forced weaning of mice. Biol Reprod 66:1471-1476 477. Travers MT, Barber MC, Tonner E, Quarrie L, Wilde CJ, Flint DJ 1996 The role of prolactin and growth hormone in the regulation of casein gene expression and mammary cell survival: relationships to milk synthesis and secretion. Endocrinology 137:1530-1539 478. Gibori G, Khan I, Warshaw ML, McLean MP, Puryear TK, Nelson S, Durkee TJ, Azhar S, Steinschneider A, Rao MC 1988 Placental-derived regulators and the complex control of luteal cell function. Recent Prog Horm Res 44:377-429 479. Soares MJ, Faria TN, Roby KF, Deb S 1991 Pregnancy and the prolactin family of hormones: coordination of anterior pituitary, uterine, and placental expression. Endocr Rev 12:402-423 480. Lahav M, Lamprecht SA, Amsterdam A, Lindner HR 1977 Suppression of 20 alpha- hydroxysteroid dehydrogenase activity in cultured rat luteal cells by prolactin. Mol Cell Endocrinol 6:293-302 481. Bjurulf E, Selstam G, Olofsson JI 1994 Increased LH receptor mRNA and extended corpus luteum function induced by prolactin and indomethacin treatment in vivo in hysterectomized pseudopregnant rats. J Reprod Fertil 102:139-145 482. Holt JA, Richards JS, Midgley AR, Jr., Reichert LE, Jr. 1976 Effect of prolactin on LH receptor in rat luteal cells. Endocrinology 98:1005-1013 483. Duan WR, Parmer TG, Albarracin CT, Zhong L, Gibori G 1997 PRAP, a prolactin receptor associated protein: its gene expression and regulation in the corpus luteum. Endocrinology 138:3216-3221 484. Manzon LA 2002 The role of prolactin in fish osmoregulation: a review. Gen Comp Endocrinol 125:291-310 485. PICKFORD GE, PHILLIPS JG 1959 Prolactin, a factor in promoting survival of hypophysectomized killifish in fresh water. Science 130:454-455 486. Ogawa M, Yagasaki M, Yamazaki F 1973 The effect of prolactin on water influx in isolated gills of the goldfish, Carassius auratus L. Comp Biochem Physiol A 44:1177- 1183 487. Maetz J, JUIEN M 1961 Action of neurohypophyseal hormones on the sodium fluxes of a freshwater teleost. Nature 189:152-153 488. Auperin B, Rentier-Delrue F, Martial JA, Prunet P 1994 Characterization of a single prolactin (PRL) receptor in tilapia (Oreochromis niloticus) which binds both PRLI and PRLII. J Mol Endocrinol 13:241-251 489. Sohm F, Pezet A, Sandra O, Prunet P, de Luze A, Edery M 1998 Activation of gene transcription by tilapia prolactin variants tiPRL188 and tiPRL177. FEBS Lett 438:119- 123 490. Sakamoto T, McCormick SD 2006 Prolactin and growth hormone in fish osmoregulation. Gen Comp Endocrinol 147:24-30 491. Pisam M, Auperin B, Prunet P, Rentier-Delrue F, Martial J, Rambourg A 1993 Effects of prolactin on alpha and beta chloride cells in the gill epithelium of the saltwater adapted tilapia "Oreochromis niloticus". Anat Rec 235:275-284

117 492. Herndon TM, McCormick SD, Bern HA 1991 Effects of prolactin on chloride cells in opercular membrane of seawater-adapted tilapia. Gen Comp Endocrinol 83:283-289 493. Peaker M, PHILLIPS JG, Wright A 1970 The effect of prolactin on the secretory activity of the nasal salt-gland of the domestic duck (Anas platyrhynchos). Journal of Endocrinology 47:123-127 494. Pippard C, Baylis PH 1986 Prolactin stimulates Na+-K+-ATPase activity located in the outer renal medulla of the rat. Journal of Endocrinology 108:95-99 495. Mainoya JR, Bern HA, Regan JW 1974 Influence of ovine prolactin on transport of fluid and sodium chloride by the mammalian intestine and gall bladder. Journal of Endocrinology 63:311-317 496. Robertson MT, Boyajian MJ, Patterson K, Robertson WV 1986 Modulation of the chloride concentration of human sweat by prolactin. Endocrinology 119:2439-2444 497. Baroni C 1967 Thymus, peripheral lymphoid tissues and immunological responsiveness of the pituitary dwarf mouse. Experientia 23:282-283 498. Baroni CD, Fabris N, Bertoli G 1969 Effects of hormones on development and function of lymphoid tissues. Synergistic action of thyroxin and somatotropic hormone in pituitary dwarf mice. Immunology 17:303-314 499. Nagy E, Berczi I, Friesen HG 1983 Regulation of immunity in rats by lactogenic and growth hormones. Acta Endocrinol (Copenh) 102:351-357 500. Berczi I, Nagy E, de Toledo SM, Matusik RJ, Friesen HG 1991 Pituitary hormones regulate c-myc and DNA synthesis in lymphoid tissue. J Immunol 146:2201-2206 501. Viselli SM, Stanek EM, Mukherjee P, Hymer WC, Mastro AM 1991 Prolactin-induced mitogenesis of lymphocytes from ovariectomized rats. Endocrinology 129:983-990 502. Hartmann DP, Holaday JW, Bernton EW 1989 Inhibition of lymphocyte proliferation by antibodies to prolactin. FASEB J 3:2194-2202 503. Tanaka T, Shiu RP, Gout PW, Beer CT, Noble RL, Friesen HG 1980 A new sensitive and specific bioassay for lactogenic hormones: measurement of prolactin and growth hormone in human serum. J Clin Endocrinol Metab 51:1058-1063 504. Murphy WJ, Durum SK, Longo DL 1993 Differential effects of growth hormone and prolactin on murine T cell development and function. J Exp Med 178:231-236 505. Russell DH, Mills KT, Talamantes FJ, Bern HA 1988 Neonatal administration of prolactin antiserum alters the developmental pattern of T- and B-lymphocytes in the thymus and spleen of BALB/c female mice. Proc Natl Acad Sci U S A 85:7404-7407 506. Murphy WJ, Rui H, Longo DL 1995 Effects of growth hormone and prolactin immune development and function. Life Sciences 57:1-14 507. Carrier M, Russell DH, Wild JC, Emery RW, Copeland JG 1987 Prolactin as a marker of rejection in human heart transplantation. J Heart Transplant 6:290-292 508. Bernton EW, Meltzer MS, Holaday JW 1988 Suppression of macrophage activation and T-lymphocyte function in hypoprolactinemic mice. Science 239:401-404 509. Woody MA, Welniak LA, Richards S, Taub DD, Tian Z, Sun R, Longo DL, Murphy WJ 1999 Use of neuroendocrine hormones to promote reconstitution after bone marrow transplantation. Neuroimmunomodulation 6:69-80 510. Clevenger CV, Sillman AL, Hanley-Hyde J, Prystowsky MB 1992 Requirement for prolactin during cell cycle regulated gene expression in cloned T-lymphocytes. Endocrinology 130:3216-3222

118 511. Dugan AL, Malarkey WB, Schwemberger S, Jauch EC, Ogle CK, Horseman ND 2004 Serum levels of prolactin, growth hormone, and cortisol in burn patients: correlations with severity of burn, serum cytokine levels, and fatality. J Burn Care Rehabil 25:306- 313 512. Krishnan N, Thellin O, Buckley DJ, Horseman ND, Buckley AR 2003 Prolactin suppresses glucocorticoid-induced thymocyte apoptosis in vivo. Endocrinology 144:2102-2110 513. BLUEM V, Fiedler K 1965 HORMONAL CONTROL REPRODUCTIVE BEHAVIOR IN SOME CICHLID FISH. Gen Comp Endocrinol 56:186-196 514. Ogawa M 1970 Effects of prolactin on the epidermal mucous cells of the goldfish, Carassius auratus L. Can J Zool 48:501-503 515. Smith RJ, Hoar WS 1967 The effects of prolactin and testosterone on the parental behaviour of the male stickleback Gasterosteus aculeatus. Anim Behav 15:342-352 516. Taniyama S, Kitahashi T, Ando H, Ban M, Ueda H, Urano A 1999 Changes in the levels of mRNAs for GH/prolactin/somatolactin family and Pit-1/GHF-1 in the pituitaries of pre-spawning chum salmon. J Mol Endocrinol 23:189-198 517. Moriya T 1982 Prolactin induces increase in the specific gravity of salamander, Hynobius retardatus, that raises adaptability to water. J Exp Zool 223:83-88 518. Janik DS, Buntin JD 1985 Behavioural and physiological effects of prolactin in incubating ring doves. Journal of Endocrinology 105:201-209 519. Youngren OM, El Halawani ME, Silsby JL, Phillips RE 1991 Intracranial prolactin perfusion induces incubation behavior in turkey hens. Biol Reprod 44:425-431 520. Crisostomo S, Guemene D, Garreau-Mills M, Morvan C, Zadworny D 1998 Prevention of incubation behavior expression in turkey hens by active immunization against prolactin. Theriogenology 50:675-690 521. March JB, Sharp PJ, Wilson PW, Sang HM 1994 Effect of active immunization against recombinant-derived chicken prolactin fusion protein on the onset of broodiness and photoinduced egg laying in bantam hens. J Reprod Fertil 101:227-233 522. Hector JA, Goldsmith AR 1985 The role of prolactin during incubation: comparative studies of three Diomedea albatrosses. Gen Comp Endocrinol 60:236-243 523. Voci VE, Carlson NR 1973 Enhancement of maternal behavior and nest building following systemic and diencephalic administration of prolactin and progesterone in the mouse. J Comp Physiol Psychol 83:388-393 524. Bridges RS, DiBiase R, Loundes DD, Doherty PC 1985 Prolactin stimulation of maternal behavior in female rats. Science 227:782-784 525. Bridges RS, Numan M, Ronsheim PM, Mann PE, Lupini CE 1990 Central prolactin infusions stimulate maternal behavior in steroid-treated, nulliparous female rats. Proc Natl Acad Sci U S A 87:8003-8007 526. Bridges RS, Mann PE 1994 Prolactin-brain interactions in the induction of material behavior in rats. Psychoneuroendocrinology 19:611-622 527. Anderson GM, Grattan DR, van den AW, Bridges RS 2006 Reproductive experience increases prolactin responsiveness in the medial preoptic area and arcuate nucleus of female rats. Endocrinology 528. Lucas BK, Ormandy CJ, Binart N, Bridges RS, Kelly PA 1998 Null mutation of the prolactin receptor gene produces a defect in maternal behavior. Endocrinology 139:4102- 4107

119 529. Prins GS, Lee C 1982 Influence of prolactin-producing pituitary grafts on the in vivo uptake, distribution, and disappearance of [3H]testosterone and [3H]dihydrotestosterone by the rat prostate lobes. Endocrinology 110:920-925 530. Prins GS, Lee C 1983 Biphasic response of the rat lateral prostate to increasing levels of serum prolactin. Biol Reprod 29:938-945 531. Prins GS 1987 Prolactin influence on cytosol and nuclear androgen receptors in the ventral, dorsal, and lateral lobes of the rat prostate. Endocrinology 120:1457-1464 532. Nevalainen MT, Valve EM, Makela SI, Blauer M, Tuohimaa PJ, Harkonen PL 1991 Estrogen and prolactin regulation of rat dorsal and lateral prostate in organ culture. Endocrinology 129:612-622 533. Ahonen TJ, Harkonen PL, Laine J, Rui H, Martikainen PM, Nevalainen MT 1999 Prolactin is a survival factor for androgen-deprived rat dorsal and lateral prostate epithelium in organ culture. Endocrinology 140:5412-5421 534. Nevalainen MT, Valve EM, Ahonen T, Yagi A, Paranko J, Harkonen PL 1997 Androgen- dependent expression of prolactin in rat prostate epithelium in vivo and in organ culture. FASEB J 11:1297-1307 535. Nevalainen MT, Valve EM, Ingleton PM, Nurmi M, Martikainen PM, Harkonen PL 1997 Prolactin and prolactin receptors are expressed and functioning in human prostate. J Clin Invest 99:618-627 536. Wennbo H, Kindblom J, Isaksson OG, Tornell J 1997 Transgenic mice overexpressing the prolactin gene develop dramatic enlargement of the prostate gland. Endocrinology 138:4410-4415 537. Robertson FG, Harris J, Naylor MJ, Oakes SR, Kindblom J, Dillner K, Wennbo H, Tornell J, Kelly PA, Green J, Ormandy CJ 2003 Prostate development and carcinogenesis in prolactin receptor knockout mice. Endocrinology 144:3196-3205 538. Nevalainen MT, Ahonen TJ, Yamashita H, Chandrashekar V, Bartke A, Grimley PM, Robinson GW, Hennighausen L, Rui H 2000 Epithelial defect in prostates of Stat5a-null mice. Lab Invest 80:993-1006 539. Buntin JD, Tesch D 1985 Effects of intracranial prolactin administration on maintenance of incubation readiness, ingestive behavior, and gonadal condition in ring doves. Horm Behav 19:188-203 540. Buntin JD, Figge GR 1988 Prolactin and growth hormone stimulate food intake in ring doves. Pharmacol Biochem Behav 31:533-540 541. Buntin JD 1989 Time course and response specificity of prolactin-induced hyperphagia in ring doves. Physiol Behav 45:903-909 542. Hnasko RM, Buntin JD 1993 Functional mapping of neural sites mediating prolactin- induced hyperphagia in doves. Brain Research 623:257-266 543. Li C, Kelly PA, Buntin JD 1995 Inhibitory effects of anti-prolactin receptor antibodies on prolactin binding in brain and prolactin-induced feeding behavior in ring doves. Neuroendocrinology 61:125-135 544. Buntin JD, Hnasko RM, Zuzick PH 1999 Role of the ventromedial hypothalamus in prolactin-induced hyperphagia in ring doves. Physiol Behav 66:255-261 545. Strader AD, Buntin JD 2001 Neuropeptide-Y: a possible mediator of prolactin-induced feeding and regulator of energy balance in the ring dove (Streptopelia risoria). J Neuroendocrinol 13:386-392

120 546. Strader AD, Buntin JD 2003 Changes in agouti-related peptide during the ring dove breeding cycle in relation to prolactin and parental hyperphagia. J Neuroendocrinol 15:1046-1053 547. Moore BJ, Gerardo-Gettens T, Horwitz BA, Stern JS 1986 Hyperprolactinemia stimulates food intake in the female rat. Brain Res Bull 17:563-569 548. Gerardo-Gettens T, Moore BJ, Stern JS, Horwitz BA 1989 Prolactin stimulates food intake in a dose-dependent manner. Am J Physiol 256:R276-R280 549. Byatt JC, Staten NR, Salsgiver WJ, Kostelc JG, Collier RJ 1993 Stimulation of food intake and weight gain in mature female rats by bovine prolactin and bovine growth hormone. Am J Physiol 264:E986-E992 550. Schwartz SM, Wade GN 1981 Effects of estradiol and progesterone on food intake, body weight, and carcass adiposity in weanling rats. Am J Physiol 240:E499-E503 551. Noel MB, Woodside B 1993 Effects of systemic and central prolactin injections on food intake, weight gain, and estrous cyclicity in female rats. Physiol Behav 54:151-154 552. Sauve D, Woodside B 1996 The effect of central administration of prolactin on food intake in virgin female rats is dose-dependent, occurs in the absence of ovarian hormones and the latency to onset varies with feeding regimen. Brain Research 729:75-81 553. Sauve D, Woodside B 2000 Neuroanatomical specificity of prolactin-induced hyperphagia in virgin female rats. Brain Research 868:306-314 554. Heil SH 1999 Sex-specific effects of prolactin on food intake by rats. Horm Behav 35:47- 54 555. Heil SH 1999 Activational and organizational actions of gonadal hormones and the sex- specific effects of prolactin on food intake by rats. Dev Psychobiol 35:61-67 556. Adler RA, Krieg RJ, Jr. 1991 Normal food intake and growth in hyperprolactinemic rats. Am J Physiol 261:R548-R552 557. Wang DY, De Stavola BL, Bulbrook RD, Allen DS, Kwa HG, Verstraeten AA, Moore JW, Fentiman IS, Chaudary M, Hayward JL, . 1987 The relationship between blood prolactin levels and risk of breast cancer in premenopausal women. Eur J Cancer Clin Oncol 23:1541-1548 558. Ferreira MF, Sobrinho LG, Pires JS, Silva ME, Santos MA, Sousa MF 1995 Endocrine and psychological evaluation of women with recent weight gain. Psychoneuroendocrinology 20:53-63 559. Greenman Y, Tordjman K, Stern N 1998 Increased body weight associated with prolactin secreting pituitary adenomas: weight loss with normalization of prolactin levels. Clin Endocrinol (Oxf) 48:547-553 560. Cohen LM, Greenberg DB, Murray GB 1984 Neuropsychiatric presentation of men with pituitary tumors (the 'four A's'). Psychosomatics 25:925-928 561. Creemers LB, Zelissen PM, 't Verlaat JW, Koppeschaar HP 1991 Prolactinoma and body weight: a retrospective study. Acta Endocrinol (Copenh) 125:392-396 562. Doknic M, Pekic S, Zarkovic M, Medic-Stojanoska M, Dieguez C, Casanueva F, Popovic V 2002 Dopaminergic tone and obesity: an insight from prolactinomas treated with bromocriptine. Eur J Endocrinol 147:77-84 563. Delgrange E, Donckier J, Maiter D 1999 Hyperprolactinaemia as a reversible cause of weight gain in male patients? Clin Endocrinol (Oxf) 50:271

121 564. Cooper GD, Pickavance LC, Wilding JP, Halford JC, Goudie AJ 2005 A parametric analysis of olanzapine-induced weight gain in female rats. Psychopharmacology (Berl) 181:80-89 565. Ota M, Mori K, Nakashima A, Kaneko YS, Fujiwara K, Itoh M, Nagasaka A, Ota A 2002 Peripheral injection of risperidone, an atypical antipsychotic, alters the bodyweight gain of rats. Clin Exp Pharmacol Physiol 29:980-989 566. Baptista T, Lacruz A, Paez X, Hernandez L, Beaulieu S 2002 The antipsychotic drug sulpiride does not affect bodyweight in male rats. Is insulin resistance involved? Eur J Pharmacol 447:91-98 567. Miller DD 2000 Review and management of clozapine side effects. J Clin Psychiatry 61 Suppl 8:14-17 568. Baptista T, Alastre T, Contreras Q, Martinez JL, Araujo dB, Paez X, Hernandez L 1997 Effects of the antipsychotic drug sulpiride on reproductive hormones in healthy men: relationship with body weight regulation. Pharmacopsychiatry 30:250-255 569. Baptista T, Lacruz A, Meza T, Contreras Q, Delgado C, Mejias MA, Hernandez L 2001 Antipsychotic drugs and obesity: is prolactin involved? Can J Psychiatry 46:829-834 570. Schotte A, Janssen PF, Gommeren W, Luyten WH, Van Gompel P, Lesage AS, De Loore K, Leysen JE 1996 Risperidone compared with new and reference antipsychotic drugs: in vitro and in vivo receptor binding. Psychopharmacology (Berl) 124:57-73 571. Ben Jonathan N, Hugo ER, Brandebourg TD, LaPensee CR 2006 Focus on prolactin as a metabolic hormone. Trends Endocrinol Metab 17:110-116 572. Brien FD 1986 Effect of suppressing prolactin in the mouse on liveweight, food intake and ovulation rate. Aust J Biol Sci 39:311-318 573. Matsuda M, Mori T, Sassa S, Sakamoto S, Park MK, Kawashima S 1996 Chronic effect of hyperprolactinemia on blood glucose and lipid levels in mice. Life Sciences 58:1171- 1177 574. Fleenor D, Oden J, Kelly PA, Mohan S, Alliouachene S, Pende M, Wentz S, Kerr J, Freemark M 2005 Roles of the lactogens and somatogens in perinatal and postnatal metabolism and growth: studies of a novel mouse model combining lactogen resistance and growth hormone deficiency. Endocrinology 146:103-112 575. Schuff KG, Hentges ST, Kelly MA, Binart N, Kelly PA, Iuvone PM, Asa SL, Low MJ 2002 Lack of prolactin receptor signaling in mice results in lactotroph proliferation and prolactinomas by dopamine-dependent and -independent mechanisms. J Clin Invest 110:973-981 576. Mercier JC, Vilotte JL 1993 Structure and function of milk protein genes. J Dairy Sci 76:3079-3098 577. Groner B, Altiok S, Meier V 1994 Hormonal regulation of transcription factor activity in mammary epithelial cells. Mol Cell Endocrinol 100:109-114 578. Kabotyanski EB, Huetter M, Xian W, Rijnkels M, Rosen JM 2006 Integration of prolactin and glucocorticoid signaling at the {beta}-casein promoter and enhancer by ordered recruitment of specific transcription factors and chromatin modifiers. Molecular Endocrinology 579. Simpson KJ, Nicholas KR 2002 The comparative biology of whey proteins. J Mammary Gland Biol Neoplasia 7:313-326

122 580. Li S, Rosen JM 1995 Nuclear factor I and mammary gland factor (STAT5) play a critical role in regulating rat whey acidic protein gene expression in transgenic mice. Mol Cell Biol 15:2063-2070 581. Oppat CA, Rillema JA 1988 Characteristics of the early effect of prolactin on lactose biosynthesis in mouse mammary gland explants. Proceedings of the Society for experimental Biology and Medicine 188:342-345 582. Vernon RG, Flint DJ 1983 Control of fatty acid synthesis in lactation. Proc Nutr Soc 42:315-331 583. Williamson DH 1990 The lactating mammary gland of the rat and the starved-refed transition: a model system for the study of the temporal regulation of substrate utilization. Biochemal Society Transections 18:853-856 584. Oftedal OT 1993 The adaptation of milk secretion to the constraints of fasting in bears, seals, and baleen whales. J Dairy Sci 76:3234-3246 585. Barber MC, Travers MT, Finley E, Flint DJ, Vernon RG 1992 Growth-hormone-prolactin interactions in the regulation of mammary and adipose-tissue acetyl-CoA carboxylase activity and gene expression in lactating rats. Biochem J 285 ( Pt 2):469-475 586. Hang J, Rillema JA 1997 Prolactin's effects on lipoprotein lipase (LPL) activity and on LPL mRNA levels in cultured mouse mammary gland explants. Proceedings of the Society for experimental Biology and Medicine 214:161-166 587. Flint DJ, Clegg RA, Vernon RG 1981 Prolactin and the regulation of adipose-tissue metabolism during lactation in rats. Mol Cell Endocrinol 22:265-275 588. Field B, Coore HG 1976 Control of rat mammary-gland pyruvate dehydrogenase by insulin and prolactin. Biochem J 156:333-337 589. Zinder O, Hamosh M, Fleck TR, Scow RO 1974 Effect of prolactin on lipoprotein lipase in mammary glands and adipose tissue of rats. Am J Physiol 226:742-748 590. Barber MC, Clegg RA, Finley E, Vernon RG, Flint DJ 1992 The role of growth hormone, prolactin and insulin-like growth factors in the regulation of rat mammary gland and adipose tissue metabolism during lactation. Journal of Endocrinology 135:195-202 591. Da Costa TH, Williamson DH 1994 Regulation of rat mammary-gland uptake of orally administered [1-14C]triolein by insulin and prolactin: evidence for bihormonal control of lipoprotein lipase activity. Biochem J 300 ( Pt 1):257-262 592. Da Costa TH, Williamson DH 1993 Effects of exogenous insulin or vanadate on disposal of dietary triacylglycerols between mammary gland and adipose tissue in the lactating rat: insulin resistance in white adipose tissue. Biochem J 290 ( Pt 2):557-561 593. Oller do Nascimento CM, Ilic V, Williamson DH 1989 Re-examination of the putative roles of insulin and prolactin in the regulation of lipid deposition and lipogenesis in vivo in mammary gland and white and brown adipose tissue of lactating rats and litter- removed rats. Biochem J 258:273-278 594. Asai-Sato M, Okamoto M, Endo M, Yoshida H, Murase M, Ikeda M, Sakakibara H, Takahashi T, Hirahara F 2006 Hypoadiponectinemia in Lean Lactating Women: Prolactin Inhibits Adiponectin Secretion from Human Adipocytes. Endocr J 595. Combs TP, Berg AH, Rajala MW, Klebanov S, Iyengar P, Jimenez-Chillaron JC, Patti ME, Klein SL, Weinstein RS, Scherer PE 2003 Sexual differentiation, pregnancy, calorie restriction, and aging affect the adipocyte-specific secretory protein adiponectin. Diabetes 52:268-276

123 596. Ling C, Billig H 2001 PRL receptor-mediated effects in female mouse adipocytes: PRL induces suppressors of cytokine signaling expression and suppresses insulin-induced leptin production in adipocytes in vitro. Endocrinology 142:4880-4890 597. Gualillo O, Lago F, Garcia M, Menendez C, Senaris R, Casanueva FF, Dieguez C 1999 Prolactin stimulates leptin secretion by rat white adipose tissue. Endocrinology 140:5149- 5153 598. Viengchareun S, Bouzinba-Segard H, Laigneau JP, Zennaro MC, Kelly PA, Bado A, Lombes M, Binart N 2004 Prolactin potentiates insulin-stimulated leptin expression and release from differentiated brown adipocytes. J Mol Endocrinol 33:679-691 599. Hamosh M, Clary TR, Chernick SS, Scow RO 1970 Lipoprotein lipase activity of adipose and mammary tissue and plasma triglyceride in pregnant and lactating rats. Biochim Biophys Acta 210:473-482 600. Smith RW 1973 The effects of pregnancy, lactation and involution on the metabolism of glucose by rat parametrial adipose tissue. J Dairy Res 40:353-360 601. Agius L, Robinson AM, Girard JR, Williamson DH 1979 Alterations in the rate of lipogenesis in vivo in maternal liver and adipose tissue on premature weaning of lactating rats: a possible regulatory role of prolactin. Biochem J 180:689-692 602. Hogan JC, Stephens JM 2005 The regulation of fatty acid synthase by STAT5A. Diabetes 54:1968-1975 603. Fortun-Lamothe L, Langin D, Lafontan M 1996 Influence of prolactin on in vivo and in vitro lipolysis in rabbits. Comp Biochem Physiol C Pharmacol Toxicol Endocrinol 115:141-147 604. Houseknecht KL, Bauman DE, Vernon RG, Byatt JC, Collier RJ 1996 Insulin-like growth factors-I and -II, somatotropin, prolactin, and placental lactogen are not acute effectors of lipolysis in ruminants. Domest Anim Endocrinol 13:239-249 605. Williams C, Coltart TM 1978 Adipose tissue metabolism in pregnancy: the lipolytic effect of human placental lactogen. Br J Obstet Gynaecol 85:43-46 606. Mochizuki M, Morikawa H, Ohga Y, Tojo S 1975 Lipolytic action of human chorionic somatomammotropin. Endocrinol Jpn 22:123-129 607. Genazzani AR, Benuzzi-Badoni M, Felber JP 1969 Human chorionic somato- mammotropine (HCSM): lipolytic action of a pure preparation on isolated fat cells. Metabolism 18:593-598 608. Hwang P, Guyda H, Friesen H 1972 Purification of human prolactin. Journal of Biological Chemistry 247:1955-1958 609. Fielder PJ, Talamantes F 1987 The lipolytic effects of mouse placental lactogen II, mouse prolactin, and mouse growth hormone on adipose tissue from virgin and pregnant mice. Endocrinology 121:493-497 610. Cabrera R, Mayor P, Calle C 1990 Effect of experimentally-induced chronic hyperprolactinemia on insulin binding and antilipolytic response in adipocytes from male rats. Rev Esp Fisiol 46:155-162 611. Ros M, Lobato MF, Garcia-Ruiz JP, Moreno FJ 1990 Integration of lipid metabolism in the mammary gland and adipose tissue by prolactin during lactation. Mol Cell Biochem 93:185-194 612. Ryan EA, Enns L 1988 Role of gestational hormones in the induction of insulin resistance. J Clin Endocrinol Metab 67:341-347

124 613. Jarrett JC, Ballejo G, Saleem TH, Tsibris JC, Spellacy WN 1984 The effect of prolactin and relaxin on insulin binding by adipocytes from pregnant women. Am J Obstet Gynecol 149:250-255 614. McAveney KM, Gimble JM, Yu-Lee L 1996 Prolactin receptor expression during adipocyte differentiation of bone marrow stroma. Endocrinology 137:5723-5726 615. Fleenor D, Arumugam R, Freemark M 2006 Growth Hormone and Prolactin Receptors in Adipogenesis: STAT-5 Activation, Suppressors of Cytokine Signaling, and Regulation of Insulin-Like Growth Factor I. Horm Res 66:101-110 616. Stewart WC, Baugh JE, Jr., Floyd ZE, Stephens JM 2004 STAT 5 activators can replace the requirement of FBS in the adipogenesis of 3T3-L1 cells. Biochem Biophys Res Commun 324:355-359 617. Nanbu-Wakao R, Morikawa Y, Matsumura I, Masuho Y, Muramatsu MA, Senba E, Wakao H 2002 Stimulation of 3T3-L1 adipogenesis by signal transducer and activator of transcription 5. Molecular Endocrinology 16:1565-1576 618. Floyd ZE, Stephens JM 2003 STAT5A promotes adipogenesis in nonprecursor cells and associates with the glucocorticoid receptor during adipocyte differentiation. Diabetes 52:308-314 619. Nanbu-Wakao R, Fujitani Y, Masuho Y, Muramatu M, Wakao H 2000 Prolactin enhances CCAAT enhancer-binding protein-beta (C/EBP beta) and peroxisome proliferator-activated receptor gamma (PPAR gamma) messenger RNA expression and stimulates adipogenic conversion of NIH-3T3 cells. Molecular Endocrinology 14:307- 316 620. Freemark M, Driscoll P, Maaskant R, Petryk A, Kelly PA 1997 Ontogenesis of prolactin receptors in the human fetus in early gestation. Implications for tissue differentiation and development. J Clin Invest 99:1107-1117 621. Freemark M, Avril I, Fleenor D, Driscoll P, Petro A, Opara E, Kendall W, Oden J, Bridges S, Binart N, Breant B, Kelly PA 2002 Targeted deletion of the PRL receptor: effects on islet development, insulin production, and glucose tolerance. Endocrinology 143:1378-1385 622. Sorenson RL, Johnson MG, Parsons JA, Sheridan JD 1987 Decreased glucose stimulation threshold, enhanced insulin secretion, and increased beta cell coupling in islets of prolactin-treated rats. Pancreas 2:283-288 623. Parsons JA, Brelje TC, Sorenson RL 1992 Adaptation of islets of Langerhans to pregnancy: increased islet cell proliferation and insulin secretion correlates with the onset of placental lactogen secretion. Endocrinology 130:1459-1466 624. Michaels RL, Sorenson RL, Parsons JA, Sheridan JD 1987 Prolactin enhances cell-to-cell communication among beta-cells in pancreatic islets. Diabetes 36:1098-1103 625. Sorenson RL, Brelje TC, Hegre OD, Marshall S, Anaya P, Sheridan JD 1987 Prolactin (in vitro) decreases the glucose stimulation threshold, enhances insulin secretion, and increases dye coupling among islet B cells. Endocrinology 121:1447-1453 626. Brelje TC, Sorenson RL 1991 Role of prolactin versus growth hormone on islet B-cell proliferation in vitro: implications for pregnancy. Endocrinology 128:45-57 627. Weinhaus AJ, Stout LE, Sorenson RL 1996 Glucokinase, hexokinase, glucose transporter 2, and glucose metabolism in islets during pregnancy and prolactin-treated islets in vitro: mechanisms for long term up-regulation of islets. Endocrinology 137:1640-1649

125 628. Petryk A, Fleenor D, Driscoll P, Freemark M 2000 Prolactin induction of insulin gene expression: the roles of glucose and glucose transporter-2. Journal of Endocrinology 164:277-286 629. Shao J, Qiao L, Friedman JE 2004 Prolactin, progesterone, and dexamethasone coordinately and adversely regulate glucokinase and cAMP/PDE cascades in MIN6 beta- cells. Am J Physiol Endocrinol Metab 286:E304-E310 630. Galsgaard ED, Nielsen JH, Moldrup A 1999 Regulation of prolactin receptor (PRLR) gene expression in insulin-producing cells. Prolactin and growth hormone activate one of the rat prlr gene promoters via STAT5a and STAT5b. Journal of Biological Chemistry 274:18686-18692 631. Brelje TC, Svensson AM, Stout LE, Bhagroo NV, Sorenson RL 2002 An immunohistochemical approach to monitor the prolactin-induced activation of the JAK2/STAT5 pathway in pancreatic islets of Langerhans. J Histochem Cytochem 50:365-383 632. Brelje TC, Stout LE, Bhagroo NV, Sorenson RL 2004 Distinctive roles for prolactin and growth hormone in the activation of signal transducer and activator of transcription 5 in pancreatic islets of langerhans. Endocrinology 145:4162-4175 633. Stout LE, Svensson AM, Sorenson RL 1997 Prolactin regulation of islet-derived INS-1 cells: characteristics and immunocytochemical analysis of STAT5 translocation. Endocrinology 138:1592-1603 634. Fleenor DE, Freemark M 2001 Prolactin induction of insulin gene transcription: roles of glucose and signal transducer and activator of transcription 5. Endocrinology 142:2805- 2810 635. Amaral ME, Ueno M, Carvalheira JB, Carneiro EM, Velloso LA, Saad MJ, Boschero AC 2003 Prolactin-signal transduction in neonatal rat pancreatic islets and interaction with the insulin-signaling pathway. Horm Metab Res 35:282-289 636. Bordin S, Amaral ME, Anhe GF, Delghingaro-Augusto V, Cunha DA, Nicoletti- Carvalho JE, Boschero AC 2004 Prolactin-modulated gene expression profiles in pancreatic islets from adult female rats. Mol Cell Endocrinol 220:41-50 637. Sorenson RL, Brelje TC, Roth C 1993 Effects of steroid and lactogenic hormones on islets of Langerhans: a new hypothesis for the role of pregnancy steroids in the adaptation of islets to pregnancy. Endocrinology 133:2227-2234 638. Weinhaus AJ, Bhagroo NV, Brelje TC, Sorenson RL 2000 Dexamethasone counteracts the effect of prolactin on islet function: implications for islet regulation in late pregnancy. Endocrinology 141:1384-1393 639. Forsyth IA 1986 Variation among species in the endocrine control of mammary growth and function: the roles of prolactin, growth hormone, and placental lactogen. J Dairy Sci 69:886-903 640. Liu Y, Hyde JF, Vore M 1992 Prolactin regulates maternal bile secretory function post partum. J Pharmacol Exp Ther 261:560-566 641. Liu Y, Suchy FJ, Silverman JA, Vore M 1997 Prolactin increases ATP-dependent taurocholate transport in canalicular plasma membrane from rat liver. Am J Physiol 272:G46-G53 642. Liu Y, Ganguly T, Hyde JF, Vore M 1995 Prolactin increases mRNA encoding Na(+)- TC cotransport polypeptide and hepatic Na(+)-TC cotransport. Am J Physiol 268:G11- G17

126 643. Ganguly TC, O'Brien ML, Karpen SJ, Hyde JF, Suchy FJ, Vore M 1997 Regulation of the rat liver sodium-dependent bile acid cotransporter gene by prolactin. Mediation of transcriptional activation by Stat5. J Clin Invest 99:2906-2914 644. Igal RA, de GD, I, Goya RG 1998 Modulation of rat liver lipid metabolism by prolactin. Prostaglandins Leukot Essent Fatty Acids 59:395-400 645. Dave JR, Brown NV, Knazek RA 1982 Prolactin modifies the prostaglandin synthesis, prolactin binding and fluidity of mouse liver membranes. Biochem Biophys Res Commun 108:193-199 646. Machida T, Taga M, Minaguchi H 1990 Effect of prolactin (PRL) on lipoprotein lipase (LPL) activity in the rat fetal liver. Asia Oceania J Obstet Gynaecol 16:261-265 647. Julve J, Robert MQ, Llobera M, Peinado-Onsurbe J 1996 Hormonal regulation of lipoprotein lipase activity from 5-day-old rat hepatocytes. Mol Cell Endocrinol 116:97- 104 648. Lorenzo M, Roncero C, Benito M 1986 The role of prolactin and progesterone in the regulation of lipogenesis in maternal and foetal rat liver in vivo and in isolated hepatocytes during the last day of gestation. Biochem J 239:135-139 649. Goodman AD, Hoekstra S, Busch RS, Meyer GS, Abend SS 1988 Effects of prolactin and growth hormone on tissue and serum carnitine in the rat. Endocrinology 123:1955- 1961 650. Zirulnik F, Anzulovich AC, Larregle E, Jahn GA, Gimenez MS 2003 Role of prolactin in the regulation of cytosolic NADP isocitrate dehydrogenase in the liver of the male rat. Endocr Res 29:201-210 651. de la Asuncion JG, Devesa A, Vina JR, Barber T 1994 Hepatic amino acid uptake is decreased in lactating rats. In vivo and in vitro studies. J Nutr 124:2163-2171 652. Franklin RB, Costello LC 1990 Prolactin directly stimulates citrate production and mitochondrial aspartate aminotransferase of prostate epithelial cells. Prostate 17:13-18 653. Franklin RB, Ekiko DB, Costello LC 1992 Prolactin stimulates transcription of aspartate aminotransferase in prostate cells. Mol Cell Endocrinol 90:27-32 654. Franklin RB, Zou J, Ma J, Costello LC 2000 Protein kinase C alpha, epsilon and AP-1 mediate prolactin regulation of mitochondrial aspartate aminotransferase expression in the rat lateral prostate. Mol Cell Endocrinol 170:153-161 655. Costello LC, Liu Y, Zou J, Franklin RB 2000 The pyruvate dehydrogenase E1 alpha gene is testosterone and prolactin regulated in prostate epithelial cells. Endocr Res 26:23-39 656. Costello LC, Liu Y, Zou J, Franklin RB 2000 Mitochondrial aconitase gene expression is regulated by testosterone and prolactin in prostate epithelial cells. Prostate 42:196-202 657. Juang HH 2004 Cyclic adenosine 3',5'-monosphosphate mediate prolactin regulation of mitochondrial aconitase in human prostate carcinoma cells. Mol Cell Endocrinol 219:141-149 658. Liu Y, Costello LC, Franklin RB 1996 Prolactin specifically regulates citrate oxidation and m-aconitase of rat prostate epithelial cells. Metabolism 45:442-449 659. Costello LC, Franklin RB 1991 Concepts of citrate production and secretion by prostate: 2. Hormonal relationships in normal and neoplastic prostate. Prostate 19:181-205

127

Chapter 2. Specific Aims of Thesis Research

128 Specific Aim I: To characterize the role of PRL in metabolic homeostasis in male and non- lactating female mice using PRL-knockouts.

Rationale: In addition to its lactogenic activity, there is considerable evidence suggesting that PRL is involved in the regulation of growth and metabolism. In rats and birds, PRL treatment significantly increases food intake and body weight. However, the information available in mice is limited and inconsistent. PRLR-deficient mice were originally reported to exhibit a reduced rate of weight gain and a striking decrease in abdominal fat. While these data suggested a significant role for PRL in metabolism, more recent studies found no differences in these animals. Moreover, it has also been reported that paradoxically, hyperprolactinemic mice have lower fat content.

Objectives: To determine the effects of PRL on 1) weight gain and body composition, 2) serum lipids and adipokines, and 3) glucose tolerance using PRL-knockout, heterozygous, and wild- type mice maintained on high-fat, low-fat, and standard chow.

Methods: Animals were weighed weekly and body composition was determined by MRI after 3 months. Serum PRL was measured using the Nb2 assay. Gene expression was determined by real time PCR. Adipokines were measured by ELISA. TG and phospholipids were measured by an enzymatic kit, while cholesterol was measured by a colorimetric assay. Blood glucose was determined using a glucometer.

Specific Aim II: To determine if PRL promotes preadipocyte proliferation and lipid accumulation.

Rationale: Adipose tissue expansion results from increases in adipocyte size, caused by lipogenesis and lipid uptake, as well as increased cell number, a result of preadipocyte proliferation and differentiation. There are multiple factors contributing to the growth of adipose tissue including dietary and hormonal factors. Because murine 3T3-L1 preadipocytes require the presence of FBS, which contains lactogenic hormones to proliferate and differentiate, PRL is hypothesized to play a role in these processes. Additionally, ectopic expression of Stat5a during adipogenesis in other preadipocyte cells increases triglyceride storage. Finally, one study using PRLR-deficient mice revealed a decrease in the number of adipocytes, further implicating PRL in preadipocyte proliferation and adipogenesis.

Objectives: To determine if PRL 1) stimulates 3T3-L1 preadipocyte proliferation, and 2) increases lipid accumulation and morphological changes during adipogenesis.

Methods: Preadipocyte proliferation was measure using the MTT, resazurin, and Calcein AM assays. Gene expression was measured using RT-PCR. Lipid accumulation was determined using Oil Red O staining and morphological changes were assessed by visual analysis.

Specific Aim III: To determine the effects of PRL on glucose uptake and lipolysis in 3T3-L1 adipocytes and adipose tissue explants.

129 Rationale: Lactogens are well-known to play an important role in glucose homeostasis during pregnancy, promoting β-cell mass and insulin production. However, the effects of PRL on glucose transport are less known. During lactation, expression of the insulin-responsive GLUT4 decreases in the mammary gland, suggesting that PRL may play a role in the regulation of glucose uptake by regulating transporter levels. By activating its receptor, PRL induces the Jak2 mediated phosphorylation of various signaling molecules, primarily the Stats, but also several IRS proteins. Therefore, PRL could exert some insulin-like effects, including glucose transport. The lipolytic actions of PRL are unclear. While some studies have shown that PRL increases glycerol release from adipocytes, others have failed to find an effect of PRL.

Objectives: 1) To assess whether PRL alters basal or insulin-stimulated rate of glucose uptake in 3T3-L1 adipocytes, 2) to determine the mechanism by which PRL exerts its effects on glucose uptake, and 3) to compare the lipolytic actions of PRL using adipose tissue explants from mice, rats, and humans.

Methods: Glucose uptake was assessed by 3H-2-deoxyglucose incorporation. Protein content was measured by the BCA assay. Real time PCR was used to quantify gene expression. For the lipolysis assay, free glycerol was measured using a colorimetric assay.

130

Chapter 3. The PRL-Deficient Mouse has an Unaltered Metabolic Phenotype

131 Published in: Endocrinology Oct;147(10):4638-45, 2006

The Prolactin-Deficient Mouse has an Unaltered Metabolic Phenotype

Christopher R. LaPensee, Nelson D. Horseman, Patrick Tso, Terry D. Brandebourg, Eric R. Hugo and Nira Ben-Jonathan

Departments of Cell Biology (CRL, TDB, ERH, NBJ), Physiology (NDH) and Pathology (PT) University of Cincinnati College of Medicine Cincinnati, OH 45267

Short Title: Metabolic parameters in the prolactin-deficient mouse

Key words: prolactin, metabolism, mouse, adiposity, adipokines, knockout

Corresponding Author: Dr. Nira Ben-Jonathan, Department of Cell Biology, University of Cincinnati, 3125 Eden Ave., Cincinnati, OH 45267-0521. Email: [email protected]

Abbreviations: CM, conditioned media; CSS, charcoal stripped serum; ELISA, enzyme-linked immunosorbent assay; FFA, free fatty acids; HF, high fat diet; LF, low fat diet; MTT, [3-(4,5- dimethylthiazol-2-yl)2,5-diphenyl tetrazolium bromide; QMR, quantitative magnetic resonance;

PRL, prolactin; PRLR, prolactin receptor; SC, standard chow.

132 Abstract

Prolactin (PRL), best recognized for its lactogenic activity, is also involved in the regulation of metabolic homeostasis in both mammalian and non-mammalian species. Although several mouse models have been used to study the metabolic functions of PRL, a clear-cut consensus has not emerged given the limited and often conflicting data. To clarify the role of PRL in metabolic homeostasis in males and non-lactating females, we used the PRL-deficient mouse. Our objectives were to compare: (1) weight gain, (2) body composition, (3) serum lipid profile, (4) circulating leptin and adiponectin levels, and (5) glucose tolerance in PRL-knockout, heterozygous, and wild-type mice maintained on standard chow, high-fat, or low-fat diets. In addition, we compared the lipolytic actions of PRL using adipose tissue explants from mice, rats and humans. We are reporting that PRL-deficiency does not affect the rate of weight gain, body composition, serum lipids or adiponectin levels in either sex on any diet. Glucose tolerance was slightly impaired in very young PRL-knockout male pups, but not in adults or in females at any age. Leptin was elevated in male, but not female, PRL-knockout mice maintained on LF diet. PRL did not affect lipolysis in adipose tissue explants from mice, but significantly inhibited glycerol release from both rat and human adipose explants in a dose-dependent manner. We conclude that PRL deficiency has negligible gross metabolic effects in mice.

133 Introduction

Prolactin (PRL) is a multifunctional, 23kDa pituitary hormone best known for its lactogenic properties. However, PRL has many physiological functions that are not limited to lactation, including osmoregulation, immune regulation and metabolism. In lower vertebrates, PRL regulates metabolic processes such as body weight and lipid content in fish, and food intake and weight gain in birds (1;2). In addition, accumulating evidence indicates that PRL is involved in several aspects of metabolic homeostasis in non-lactating female and male mammals (3).

PRL has well-defined metabolic activities in target tissues such the mammary gland, where it stimulates the synthesis of milk proteins, lactose and lipids (4), the pancreas, where it promotes β-cell growth and insulin production (5;6), and the prostate, where it increases citrate biosynthesis (7). There is also an emerging recognition that PRL directly regulates adipose tissue function. Several isoforms of the PRLR are expressed in white and brown adipose tissue of a number of species (8-12). Studies with primary adipocytes, cell lines, or adipose tissue explants demonstrate that PRL suppresses lipogenesis by down-regulating lipoprotein lipase and fatty acid synthase (10;13;14), whereas its lipolytic activity varies across species (15-17).

PRL also regulates the production and secretion of adipokines. Adipose tissue is a dynamic organ, acting both as an energy storage depot and as a hormone secreting endocrine tissue. Adipokines affect metabolic homeostasis by acting on the brain, liver, pancreas and muscle (18;19). Recent reports indicate that PRL inhibits the production of interleukin-6 (8) and adiponectin (20;21), both of which are involved in obesity-related insulin resistance. On the other hand, the relationship between PRL and leptin, a satiety factor which regulates food intake and energy expenditure, is less clear (22-25).

Body weight and adiposity are coordinated in a complex manner by nutritional factors, hormonal signals and energy expenditure. Information on the involvement of PRL with these parameters is primarily based on observations made in rats, whereby long-term elevation of circulating PRL results in increased body weight with inconsistent changes in adipose tissue mass (26-29). In humans, chronic hyperprolactinemia is sometimes associated with weight gain, which can be reversed upon treatment with bromocriptine, which normalizes serum PRL levels (30;31). However, this weight loss does not occur in all patients, is modest and delayed, and does not correlate well with the rapid suppression of serum PRL. Moreover, bromocriptine may have independent effects on weight gain and adiposity.

Mice are used extensively to study metabolic homeostasis, given the ease of generating transgenics overexpressing or lacking specific genes. Despite the use of several mouse models to examine the metabolic functions of PRL, limited and often conflicting data are available. In PRL-overexpressing females, a minor decrease in adiposity was observed with no difference in body weight (9). A 2001 study, using the PRL receptor (PRLR)-deficient mouse, reported a significant reduction in the rate of weight gain and a marked decrease in abdominal fat mass (22), while a more recent study found no difference in these animals (32). Paradoxically, male mice with chronic hyperprolactinemia have lower epididymal fat content (33).

134 To clarify the metabolic actions of PRL we used the PRL-deficient mouse (34). A major advantage of this experimental model is that the PRLR is expressed and functional, enabling the application of replacement therapy aimed at reversing any metabolic alterations caused by PRL deficiency. Our objectives were to compare: a) weight gain, b) body composition, c) serum lipid profile, d) circulating leptin and adiponectin concentrations, and e) glucose tolerance in PRL- knockout, heterozygous, and wild-type mice fed standard chow (SC), high-fat (HF), or low-fat (LF) diets. We also examined whether PRL deficiency altered PRLR expression in adipose tissue and liver. In addition, we compared the lipolytic activity of PRL using adipose tissue explants from wild-type mice, PRL-knockout mice, rats, and humans.

Materials and methods

Animals

The generation of PRL knockout mice has been described previously (34). Wild-type (+/+), heterozygous (+/-), and PRL knockout (-/-) mice were generated by breeding +/- or -/- males with +/- females. Pups were genotyped by PCR analysis of DNA isolated from tail clippings. Primers targeted to the 5’ sequence of the neomycin resistance gene (PKG-neo), inserted in the coding region of the PRL protein, amplified an expected product in +/- mice and -/- mice. Primers for the 5’ sequence of the PRL gene, upstream of the neo insert, yielded an expected product in +/+ and +/- mice, but not in -/- mice. A common 3’ primer for exon 4 of the PRL gene was used to amplify both products. Adult male rats (Sprague Dawley) were obtained from Harlan (Indianapolis, Indiana). Mice and rats were housed on a 12-h light, 12-h dark light cycle (lights on at 0600h), with food and water available ad libitum unless otherwise specified. Animals were fasted overnight before sacrificed by CO2 inhalation and cervical dislocation between 9-11 am. Blood was collected by cardiac puncture and centrifuged at 5,000xg for 10 minutes at 4°C. Serum was kept at -80 C for later analysis. Liver and adipose tissue (epididymal or peri-uterine) were removed and frozen for PRLR analysis by RT-PCR. Animal protocols were approved by the University of Cincinnati Institutional Animal Care and Use Committee.

Human patients

Fresh subcutaneous abdominal adipose tissue was obtained from patients undergoing abdominoplasty. The study was approved by the University of Cincinnati Institutional Review Board and informed consent was obtained from each patient.

NB2 bioassay for PRL

Rat Nb2 lymphocytes were cultured as previously described (35). Briefly, cells were plated in 96-well plates (20,000 cells/well) and incubated with mouse PRL (mPRL) (NIDDK) in triplicate and mouse serum in duplicate (0.5, 1, and 2 μl). After three days, cell number was determined by the MTT method. The amount of PRL in the serum was calculated from the standard curve, with a lowest detectable level of 0.2 ng/ml.

Real-Time PCR

135 Total RNA was extracted using Tri Reagent (MRC, Cincinnati, OH), and 5 μg of RNA was reverse transcribed using Superscript II reverse transcriptase (Invitrogen, Carlsbad, CA) as previously described (8). Real-time PCR was performed on 200 ng of cDNA using intron- spanning primers for mPRLR (forward 5’- CAC AGT AAA TGC CAC GAA CG - 3’, reverse 5’ - GGC AAC CAT TTT ACC CAC AG - 3’) and β-Actin as a control (forward 5’- ACT GCT CTG GCT CCT AGC AC - 3’, reverse 5’ - AGT CCG CCT AGA AGC ACT TG - 3’). The PRLR primers (located in the extracellular domain of the receptor) were designed so as to detect all isoforms. Products were amplified on a SmartCycler I (Cepheid, Sunnyvale, CA) using Immolase heat-activated Taq DNA polymerase (Bioline, Randolph MA) and SYBR Green I (Invitrogen Life Technologies, Inc.) for fluorometric product detection. Cycle parameters were 96 C for 6 min for polymerase activation, followed by 40 cycles of 95 C for 15 sec, 57 C for 15 sec, and 72 C for 25 sec, and optical read stage at 83.5 C for 6 sec. Product purity was confirmed by DNA melting curve analysis. Fold changes in gene expression were calculated from the cycle threshold measurements, using the method of Pfaffl et al (36), and data were expressed as fold induction vs wild type mice.

Diets

Mice at 28 days of age were randomly divided into 3 experimental diet groups. Standard chow (SC), obtained from Harlan, was comprised of 10% fat, 20% protein, and 3% fiber. High-fat (HF) and low-fat (LF) diets were prepared by Dyets Inc., Bethlehem, PA. The HF diet (20% fat) contained 20g of fat/100g of food (19g butter fat and 1g soybean oil to provide essential fatty acids). The LF diet contained 4% fat (3 g butter fat and 1 g soybean oil). Protein, essential minerals, and vitamins were equal in the HF and LF diets. Animals were maintained on the specific diets for at least 3 months and were weighed weekly.

Body Composition

Quantitative magnetic resonance (QMR, Echo Medical Systems, Houston TX) was used for in vivo body composition analysis. Mice from each diet group were analyzed at 14 weeks of age. The percent lean and fat body mass were calculated by dividing tissue mass by total body weight.

Serum Lipid Profile

Triglycerides were measured using a kit from Randox (Crumlin, UK). The Wako enzymatic method was used to measure phospholipids and non-esterified fatty acids (37). Cholesterol was measured by a colorimetric assay from Thermo Electron Corporation (Waltham, MA, USA).

Leptin and Adiponectin Determination

Serum leptin and adiponectin were measured using ELISA kits purchased from R&D Systems (Minneapolis, MN). The lowest level of detection was 62.5 pg/ml for leptin, and 160 pg/ml for adiponectin.

Glucose Tolerance Test

136

Glucose tolerance was measured at 4, 8, and 12 weeks of age. Prior to the test, animals were fasted overnight. Tail blood was obtained 15 minutes before, and 15, 30, 60 and 120 min after injection of D-glucose (10% in water, 10 µl/g body weight). Blood was directly analyzed for glucose using a One Touch glucometer (Lifescan, Milpitas, CA).

Lipolysis

Rats or mice were fasted overnight before the experiment. Epididymal fat was removed, minced and blood vessels carefully dissected out. Subcutaneous abdominal fat was obtained from patients undergoing abdominoplasty. In each case, explants (4-5 pieces totaling ∼50 mg) were incubated in Medium 199 (Cellgro, Herndon VA) containing, 1% charcoal stripped serum (CSS) (Hyclone, Logan UT). Samples were incubated with recombinant ovine PRL (for mice or rats) or human PRL (for human explants) at 0, 1, 5, and 25 ng/ml for 24 hrs. For basal lipolysis, explants were incubated in 200µl KRH (Krebs-Ringers-Hepes, 1.8 mM CaCl, 1 mM sodium pyruvate, 1% CSS) including the same PRL doses as above for 2 hrs. Conditioned media (CM) were collected and replaced with the same medium containing 100nM isoproterenol for 2 hrs. CM were again collected and glycerol release was measured by a colorimetric assay (Sigma). Glycerol release was calculated from a standard curve and normalized by tissue weight. Data are expressed as nmol glycerol/mg tissue/2 hrs. Data analysis

Statistical differences were determined by one-way ANOVA followed by Fisher LSD post hoc analysis. For glucose tolerance, the area under the curve was calculated by GraphPAD PRIZM. All experiments were repeated at least three times. P values <0.05 were considered significant.

Results

Serum PRL levels in the three mouse genotypes

The Nb2 bioassay was used to measure serum PRL levels in males and randomly cycling females. As shown in Fig 1, left panel, Nb2 cell number showed a curvilinear relationship with the amount of mPRL from 0-75 pg/well. Addition of 0.5, 1, and 2 μl of serum from +/+ males showed parallelism with the linear portion of the curve. A similar dose-response relationship was observed using serum from +/- mice (not shown). Serum from -/- mice was below the lower limit of detection in all aliquots tested. Serum PRL concentrations were higher in females than males (table inset), with no apparent differences between +/- and +/+ of either sex. Fig 1, right panel shows that PRLR expression in adipose tissue and liver was unchanged in all female genotypes. Similar results were obtained in liver and epididymal adipose tissue of males (not shown).

Body weight and composition

To compare the growth rate of the three genotypes, animals were placed under different diet paradigms and weighed weekly. Weaning weights of both PRL-/- males and females were comparable to +/+ and +/- littermates. There was no difference in growth rates between the three genotypes of either females (Fig 2A) or males (Fig 2B) on SC. Animals on LF diet gained weight

137 at a slower rate than those fed SC, with no apparent differences between the three genotypes. As expected, mice on HF diet gained considerably more (~20%) weight than those on LF diet, but again, there were no differences in body weight between the three genotypes of either males or females. PRL-/- males on HF diet showed a tendency for a heavier weight than +/- or +/+ mice, but the difference was not statistically significant. QMR was used to compare body composition in the HF and LF experimental groups at 14 weeks of age. As expected, both males and females on HF diet accumulated approximately twice as much body fat as those on LF diet (Fig 3, left panels). However, no differences in the percent of body fat were evident between the three genotypes maintained on either diet. PRL-/- females on HF diet showed a tendency for a higher body fat content but the difference was significant only compared to +/- females. No significant changes in lean tissue mass were observed among all experimental groups (Fig 3, right panels). A group of male mice on LF diet was subjected to body composition analysis at 4, 8, and 12 weeks of age. However, there were no differences in either fat or lean tissue mass between the three genotypes at any time (data not shown).

Serum lipid profile

Serum from males maintained on HF, LF or SC diets for 12-14 weeks was analyzed for triglycerides, cholesterol, phospholipids, and non-esterified free fatty acids (FFA). As shown in Fig 4, there were no differences in these parameters among the three genotypes on any diet. HF food increased cholesterol levels as compared to SC and LF food, but without significant differences among the three genotypes. Analysis of serum from females also showed no differences, except for decreased FFA in -/- mice fed LF food (data not shown).

Elevated leptin in male mice on LF diet

Serum from mice maintained on HF and LF diets for 12-14 weeks were analyzed for leptin and adiponectin. As shown in Fig 5, left panel, leptin concentration was significantly higher (6.2±0.7 ng/ml) in -/- males on LF diet compared to +/- (3.8±0.7 ng/ml) and +/+ (4.0±.6 ng/ml) mice. No differences were observed in females on LF diet (data not shown). As expected, leptin was significantly higher in mice on HF diet, but no differences were observed among the three genotypes in either males or females. Adiponectin levels (Fig 5, right panel) were unchanged between -/- and +/+ males on either diet. Females had a 50% higher serum adiponectin levels than males, but also showed no difference among the genotypes (data not shown).

Impaired glucose tolerance response only in 4-week old male mice

Males were subjected to glucose tolerance tests at 4, 8, and 12 weeks of age. At all ages, fasted blood glucose levels were similar in the three genotypes. In 4-week old mice, peak glucose levels after ip glucose injection were comparable in all genotypes (Fig 6, left panel). However, the total area under the curve in -/- mice was significantly (p<0.05) higher than that in +/+ mice. By 8 and 12 weeks of age, there were no differences in glucose tolerance among the three genotypes (Fig 6, middle and right panels). At 4 and 8 weeks of age, blood glucose levels after ip glucose injection were similar in all female genotypes (data not shown)

PRL inhibits lipolysis in adipose explants from rats and humans but not mice

138

The effect of PRL on lipolysis was compared in epididymal adipose tissue explants obtained from mice and rats, and subcutaneous explants from humans. As evident in Fig. 7, upper panels, PRL did not affect basal or isoproterenol-stimulated lipolysis in either +/+ or -/- mice. On the other hand, PRL at physiological concentrations caused a significant, dose-dependent inhibition of both basal and isoproterenol-stimulated lipolysis in adipose explants from rats and humans (Fig 7, lower panels). In all cases, isoproterenol-stimulated glycerol release was 2-4 fold above basal release.

Discussion

We are reporting that PRL-deficiency in mice does not alter the rate of weight gain, body composition, serum lipids or circulating adiponectin levels. There are no differences in PRLR expression in either liver or adipose tissue among the three genotypes. Glucose tolerance is impaired in very young PRL-knockout males but not in adult males or females at any age. Serum leptin levels are elevated in male, but not female, PRL-knockout mice maintained on LF diet. PRL does not affect lipolysis in adipose tissue explants from mice but has a significant, dose- dependent inhibition of lipolysis in explants from rats and humans. These comprehensive studies indicate that although PRL-deficiency has minor effects on leptin release and glucose tolerance, it does not result in global changes in weight gain and adiposity in mice of either sex. The suppression of lipolysis by PRL in adipose tissue from rats and humans, but not mice, gives further credence to the notion that the metabolic effects of PRL are species-specific.

Utilizing the very sensitive Nb2 bioassay, we confirm that PRL is undetectable in serum from -/- male or female mice. As expected, serum PRL levels are higher in wild type or heterozygous females than males. The similarity of circulating PRL levels in +/+ and +/- mice of either sex indicates that loss of one PRL allele does not affect overall PRL production/release. However, because PRL levels may differ between wild-type and heterozygous females during the estrous cycle or under stress in either males or females, we included heterozygotes in all our studies. Notably, all of our studies were conducted during the first five months of life, prior to the formation of pituitary tumors. These develop with aging in both PRL- and PRLR-deficient mice (38;39), and likely result in altered pituitary function.

Since the metabolic status of an organism depends on its nutritional as well as hormonal status, mice were subjected to several diet paradigms. For the longitudinal analysis of growth rates we used a large number of mice of both sexes to ensure the significance of any observations. PRL- deficiency did not affect weight gain in males or females under any diet paradigm (Fig 2). As revealed by QMR, a precise tool for measuring total fat within an intact organism, PRL deficiency did not alter fat content in these mice (Fig 3). Our data conflict with the report on reduced weight gain in PRLR-deficient mice (22), which was not reproduced in later studies by the same authors (32;40). Similarly, our results do not agree with the report that both male and female PRLR-deficient mice had a marked reduction in abdominal fat mass at 8-9 months of age, as determined by gross dissection of abdominal fat (22). Indeed, a subsequent study, using dual- energy x-ray absorptiometry (DEXA), revealed no difference in fat content of either 8-12 month old males or 4 month old females (32). Notably, some studies with the PRLR-deficient mouse did not take into account the confounding influence of pituitary tumors in older animals.

139

Studies from other laboratories also failed to find a clear relationship between PRL, weight gain and adiposity in mice. For example, minor changes in weight gain and fat content were seen in mice made hyperprolactinemic by PRL overexpression or pituitary grafts (9;33). Suppression of PRL with bromocriptine in ob/ob male mice caused no changes in body weight, with slight reductions in food intake and hyperglycemia (41). In contrast to mice, chronic elevation of PRL in rats has consistently been linked to increased food intake and weight gain, while PRL suppression resulted in the opposite outcome (26;27;42;43). Sustained hyperprolactinemia in humans, caused by pituitary tumors or treatment with antipsychotic drugs, is occasionally accompanied by increased weight (30;31;44;45), but it is unclear whether the elevated PRL is causative or coincidental to the weight gain.

During lactation, PRL acts as a physiological sensor that responds to the demands for milk production by partitioning lipids away from adipose tissue in favor of the mammary gland (3). In lactating animals, PRL suppresses lipoprotein lipase (LPL) activity in adipose tissue while increasing its activity in the mammary gland (13;46). Since LPL is the major enzyme that hydrolyzes circulating lipoprotein-triglyceride complexes, changes in its activity should have an impact on serum lipid profile. Nonetheless, it is unclear whether PRL alters lipid metabolism in non-lactating mice, with one study reporting a small reduction in serum FFA in males receiving pituitary grafts (33). Our results revealed no noticeable differences in triglycerides, cholesterol, phospholipids, or FFAs between the three genotypes in males or females on either diet (Fig 4).

We observed a significant, though modest, increase in serum leptin levels in male PRL-/- mice on LF diet but not in those on HF diet (Fig 5). Increased production of leptin driven by the HF diet-induced adiposity may have been sufficiently high to override any discernible effects of PRL-deficiency. Unlike males, there was no difference in serum leptin levels between +/+ and -/- females on either diet. Our data do not support the report on lower plasma leptin levels in PRLR- deficient females (22), but agree with a later report showing no difference in leptin levels in such mice (32). Leptin secretion is differentially regulated in males and females, with estrogens stimulating and androgens inhibiting its production (47). It remains to be determined whether PRL interacts with the gonadal steroids in the regulation of leptin release. The reports of a direct effect of PRL on leptin production by adipocytes are also conflicting. In isolated mouse adipocytes, PRL alone has no effect on leptin, but inhibits insulin-induced leptin production (24). In contrast, treatment with PRL and insulin increases leptin release from T37i brown adipocytes (25). These discrepancies may be due to differences in experimental design, doses of PRL or depot-specific control of leptin release.

Adiponectin production is higher in females than males, and is negatively correlated with adiposity (48). An inhibitory effect of PRL on adiponectin release in female mice has been reported in two recent studies. In normal females, infusion of PRL suppressed circulating adiponectin whereas bromocriptine injection caused an elevation (20). Serum adiponectin was significantly lower in PRL-overexpressing females, but not males; no changes were seen in male or female PRLR-deficient mice (21). PRL also inhibited adiponectin release from subcutaneous abdominal adipose explants from healthy, non-obese women (21). Our studies reveal no difference in serum adiponectin levels between PRL-/- or +/+ mice of either sex (Fig 5). Interestingly, neither the nutritional status nor adiposity altered circulating adiponectin levels,

140 although adiponectin is known to be reduced in obese individuals. Nilsson et al. suggested that adiponectin levels are more reflective of insulin sensitivity rather than adiposity (21).

It is well established that lactogens promote β-cell growth and insulin production (6;49;50). Indeed, PRLR-deficiency results in reduced β-cell mass and islet density and impaired glucose tolerance in adult male and female mice (40). In our case, the clearance of blood glucose following glucose injection was delayed in 4-week old PRL-/- males (Fig 6), but this impairment did not extend to older animals. We do not know whether the transient reduction of glucose tolerance was due to a delayed maturation of pancreatic function or to a lower insulin sensitivity, and if so, why the female PRL-/- pups or adult mice did not show this impairment. Since the PRLR is expressed in the pancreas in late gestation (11), exposure to placental lactogens in utero could have supported β-cell development in our animals while the PRLR-null mice are unable to respond to either PRL or placental lactogens.

An older study reported that PRL induced lipolysis in adipose tissue explants from virgin or pregnant mice at an extremely high dose of 5 μg/ml (15). In the present studies, when used within the physiological range (1, 5, and 25 ng/ml), PRL did not affect glycerol release from mouse epididymal adipose explants from either -/- or +/+ mice (Fig 7). PRL treatment at a concentration as high as 125 ng/ml also had no effect on lipolysis in mouse adipose tissue (data not shown). In contrast, treatment with low doses of PRL inhibited basal- and isoproterenol- stimulated lipolysis in rat epididymal explants by as much as 45%, with a lesser, though significant, inhibitory effect on lipolysis in subcutaneous adipose explants obtained from women. Similar results were observed in samples obtained from men (data not shown). The ligand used to treat mouse and rat explants was ovine PRL, which is active in adipocytes from both species (14;23). Since a 2 hr incubation with PRL does not affect lipolysis in rat epididymal explants (data not shown), we speculate that PRL may act by downregulating hormone-sensitive lipase or adipose triglyceride lipase, the critical enzymes responsible for catabolism of stored triglycerides (51;52), rather than by affecting their phosphorylation or that of perilipin, as is the case with catecholamines (53). The mechanism by which PRL inhibits lipolysis is currently under investigation.

In conclusion, PRL is a multifunctional hormone which plays modulating and adaptive roles in many physiological systems across the animal kingdom. Among the several hundred functions ascribed to PRL, not all are equally important in any one species. For example, the osmoregulatory role of PRL is best exemplified in fish, its effects on hair follicles are more pronounced in sheep, while the control of corpus luteum function by PRL is unique to rodents. It is not totally surprising, therefore, that PRL appears to have gross metabolic effects in rats but not in mice. Similar to mice, there is no strong evidence to suggest that PRL is an important factor in the regulation of body weight in humans. However, PRL may play a role in other aspects of metabolic homeostasis such as lipolysis and adipokine release. Although our studies yielded mostly negative data, they are important for several reasons. First, they add to the fund of knowledge on the comparative aspects of PRL. Second, they address many published controversies with respect to the metabolic activity of PRL in the mouse. And finally, they should serve as a guidance for investigators wishing to study the roles of PRL in metabolic homeostasis in their selection of the most appropriate species.

141 Acknowledgements

This work was supported by NIH grants ES012212 and CA096613, DOD BC05725 and Susan G. Komen Breast Cancer Foundation grant BCRT87406 (to N.B.J.), and NIH grant DK52134 (to NDH).

References

1. Meier AH 1969 Diurnal variations of metabolic responses to prolactin in lower vertebrates. Gen Comp Endocrinol 12:55-62 2. Buntin JD, Hnasko RM, Zuzick PH 1999 Role of the ventromedial hypothalamus in prolactin-induced hyperphagia in ring doves. Physiol Behav 66:255-261 3. Ben Jonathan N, Hugo ER, Brandebourg TD, Lapensee CR 2006 Focus on prolactin as a metabolic hormone. Trends Endocrinol Metab 4. Neville MC, McFadden TB, Forsyth I 2002 Hormonal regulation of mammary differentiation and milk secretion. J Mammary Gland Biol Neoplasia 7:49-66 5. Sorenson RL, Brelje TC 1997 Adaptation of islets of Langerhans to pregnancy: beta-cell growth, enhanced insulin secretion and the role of lactogenic hormones. Horm Metab Res 29:301-307 6. Sorenson RL, Brelje TC, Hegre OD, Marshall S, Anaya P, Sheridan JD 1987 Prolactin (in vitro) decreases the glucose stimulation threshold, enhances insulin secretion, and increases dye coupling among islet B cells. Endocrinology 121:1447-1453 7. Costello LC, Franklin RB 2002 Testosterone and prolactin regulation of metabolic genes and citrate metabolism of prostate epithelial cells. Horm Metab Res 34:417-424 8. Hugo ER, Brandebourg TD, Comstock CE, Gersin KS, Sussman JJ, Ben Jonathan N 2006 LS14: a novel human adipocyte cell line that produces prolactin. Endocrinology 147:306-313 9. Ling C, Hellgren G, Gebre-Medhin M, Dillner K, Wennbo H, Carlsson B, Billig H 2000 Prolactin (PRL) receptor gene expression in mouse adipose tissue: increases during lactation and in PRL-transgenic mice. Endocrinology 141:3564-3572 10. Ling C, Svensson L, Oden B, Weijdegard B, Eden B, Eden S, Billig H 2003 Identification of functional prolactin (PRL) receptor gene expression: PRL inhibits lipoprotein lipase activity in human white adipose tissue. J Clin Endocrinol Metab 88:1804-1808 11. Royster M, Driscoll P, Kelly PA, Freemark M 1995 The prolactin receptor in the fetal rat: cellular localization of messenger ribonucleic acid, immunoreactive protein, and ligand- binding activity and induction of expression in late gestation. Endocrinology 136:3892- 3900 12. Symonds ME, Phillips ID, Anthony RV, Owens JA, McMillen IC 1998 Prolactin receptor gene expression and foetal adipose tissue. J Neuroendocrinol 10:885-890 13. Flint DJ, Binart N, Kopchick J, Kelly P 2003 Effects of growth hormone and prolactin on adipose tissue development and function. Pituitary 6:97-102 14. Hogan JC, Stephens JM 2005 The regulation of fatty acid synthase by STAT5A. Diabetes 54:1968-1975

142 15. Fielder PJ, Talamantes F 1987 The lipolytic effects of mouse placental lactogen II, mouse prolactin, and mouse growth hormone on adipose tissue from virgin and pregnant mice. Endocrinology 121:493-497 16. Houseknecht KL, Bauman DE, Vernon RG, Byatt JC, Collier RJ 1996 Insulin-like growth factors-I and -II, somatotropin, prolactin, and placental lactogen are not acute effectors of lipolysis in ruminants. Domest Anim Endocrinol 13:239-249 17. Fortun-Lamothe L, Langin D, Lafontan M 1996 Influence of prolactin on in vivo and in vitro lipolysis in rabbits. Comp Biochem Physiol C Pharmacol Toxicol Endocrinol 115:141-147 18. Ahima RS, Saper CB, Flier JS, Elmquist JK 2000 Leptin regulation of neuroendocrine systems. Front Neuroendocrinol 21:263-307 19. Ahima RS, Flier JS 2000 Adipose tissue as an endocrine organ. Trends Endocrinol Metab 11:327-332 20. Combs TP, Berg AH, Rajala MW, Klebanov S, Iyengar P, Jimenez-Chillaron JC, Patti ME, Klein SL, Weinstein RS, Scherer PE 2003 Sexual differentiation, pregnancy, calorie restriction, and aging affect the adipocyte-specific secretory protein adiponectin. Diabetes 52:268-276 21. Nilsson L, Binart N, Bohlooly Y, Bramnert M, Egecioglu E, Kindblom J, Kelly PA, Kopchick JJ, Ormandy CJ, Ling C, Billig H 2005 Prolactin and growth hormone regulate adiponectin secretion and receptor expression in adipose tissue. Biochem Biophys Res Commun 331:1120-1126 22. Freemark M, Fleenor D, Driscoll P, Binart N, Kelly P 2001 Body weight and fat deposition in prolactin receptor-deficient mice. Endocrinology 142:532-537 23. Gualillo O, Lago F, Garcia M, Menendez C, Senaris R, Casanueva FF, Dieguez C 1999 Prolactin stimulates leptin secretion by rat white adipose tissue. Endocrinology 140:5149- 5153 24. Ling C, Billig H 2001 PRL receptor-mediated effects in female mouse adipocytes: PRL induces suppressors of cytokine signaling expression and suppresses insulin-induced leptin production in adipocytes in vitro. Endocrinology 142:4880-4890 25. Viengchareun S, Bouzinba-Segard H, Laigneau JP, Zennaro MC, Kelly PA, Bado A, Lombes M, Binart N 2004 Prolactin potentiates insulin-stimulated leptin expression and release from differentiated brown adipocytes. J Mol Endocrinol 33:679-691 26. Baptista T, de Baptista EA, Lalonde J, Plamondon J, Kin NM, Beaulieu S, Joober R, Richard D 2004 Comparative effects of the antipsychotics sulpiride and risperidone in female rats on energy balance, body composition, fat morphology and macronutrient selection. Prog Neuropsychopharmacol Biol Psychiatry 28:1305-1311 27. Byatt JC, Staten NR, Salsgiver WJ, Kostelc JG, Collier RJ 1993 Stimulation of food intake and weight gain in mature female rats by bovine prolactin and bovine growth hormone. Am J Physiol 264:E986-E992 28. Gerardo-Gettens T, Moore BJ, Stern JS, Horwitz BA 1989 Prolactin stimulates food intake in a dose-dependent manner. Am J Physiol 256:R276-R280 29. Sauve D, Woodside B 2000 Neuroanatomical specificity of prolactin-induced hyperphagia in virgin female rats. Brain Res 868:306-314 30. Doknic M, Pekic S, Zarkovic M, Medic-Stojanoska M, Dieguez C, Casanueva F, Popovic V 2002 Dopaminergic tone and obesity: an insight from prolactinomas treated with bromocriptine. Eur J Endocrinol 147:77-84

143 31. Greenman Y, Tordjman K, Stern N 1998 Increased body weight associated with prolactin secreting pituitary adenomas: weight loss with normalization of prolactin levels. Clin Endocrinol (Oxf) 48:547-553 32. Fleenor D, Oden J, Kelly PA, Mohan S, Alliouachene S, Pende M, Wentz S, Kerr J, Freemark M 2005 Roles of the lactogens and somatogens in perinatal and postnatal metabolism and growth: studies of a novel mouse model combining lactogen resistance and growth hormone deficiency. Endocrinology 146:103-112 33. Matsuda M, Mori T, Sassa S, Sakamoto S, Park MK, Kawashima S 1996 Chronic effect of hyperprolactinemia on blood glucose and lipid levels in mice. Life Sci 58:1171-1177 34. Horseman ND, Zhao W, Montecino-Rodriguez E, Tanaka M, Nakashima K, Engle SJ, Smith F, Markoff E, Dorshkind K 1997 Defective mammopoiesis, but normal hematopoiesis, in mice with a targeted disruption of the prolactin gene. EMBO J 16:6926-6935 35. Zinger M, McFarland M, Ben-Jonathan N 2003 Prolactin expression and secretion by human breast glandular and adipose tissue. Journal of Clinical Endocrinology and Metabolism 88:689-696 36. Pfaffl MW, Horgan GW, Dempfle L 2002 Relative expression software tool (REST) for group-wise comparison and statistical analysis of relative expression results in real-time PCR. Nucleic Acids Res 30:e36 37. Takayama M, Itoh S, Nagasaki T, Tanimizu I 1977 A new enzymatic method for determination of serum choline-containing phospholipids. Clin Chim Acta 79:93-98 38. Cruz-Soto ME, Scheiber MD, Gregerson KA, Boivin GP, Horseman ND 2002 Pituitary tumorigenesis in prolactin gene-disrupted mice. Endocrinology 143:4429-4436 39. Schuff KG, Hentges ST, Kelly MA, Binart N, Kelly PA, Iuvone PM, Asa SL, Low MJ 2002 Lack of prolactin receptor signaling in mice results in lactotroph proliferation and prolactinomas by dopamine-dependent and -independent mechanisms. J Clin Invest 110:973-981 40. Freemark M, Avril I, Fleenor D, Driscoll P, Petro A, Opara E, Kendall W, Oden J, Bridges S, Binart N, Breant B, Kelly PA 2002 Targeted deletion of the PRL receptor: effects on islet development, insulin production, and glucose tolerance. Endocrinology 143:1378-1385 41. Cincotta AH, Tozzo E, Scislowski PW 1997 Bromocriptine/SKF38393 treatment ameliorates obesity and associated metabolic dysfunctions in obese (ob/ob) mice. Life Sci 61:951-956 42. Moore BJ, Gerardo-Gettens T, Horwitz BA, Stern JS 1986 Hyperprolactinemia stimulates food intake in the female rat. Brain Res Bull 17:563-569 43. Noel MB, Woodside B 1993 Effects of systemic and central prolactin injections on food intake, weight gain, and estrous cyclicity in female rats. Physiol Behav 54:151-154 44. Kopelman PG 2000 Physiopathology of prolactin secretion in obesity. Int J Obes Relat Metab Disord 24 Suppl 2:S104-S108 45. Baptista T, Lacruz A, Meza T, Contreras Q, Delgado C, Mejias MA, Hernandez L 2001 Antipsychotic drugs and obesity: is prolactin involved? Can J Psychiatry 46:829-834 46. Ros M, Lobato MF, Garcia-Ruiz JP, Moreno FJ 1990 Integration of lipid metabolism in the mammary gland and adipose tissue by prolactin during lactation. Mol Cell Biochem 93:185-194

144 47. Mystkowski P, Schwartz MW 2000 Gonadal steroids and energy homeostasis in the leptin era. Nutrition 16:937-946 48. Berg AH, Combs TP, Scherer PE 2002 ACRP30/adiponectin: an adipokine regulating glucose and lipid metabolism. Trends Endocrinol Metab 13:84-89 49. Brelje TC, Scharp DW, Lacy PE, Ogren L, Talamantes F, Robertson M, Friesen HG, Sorenson RL 1993 Effect of homologous placental lactogens, prolactins, and growth hormones on islet B-cell division and insulin secretion in rat, mouse, and human islets: implication for placental lactogen regulation of islet function during pregnancy. Endocrinology 132:879-887 50. Brelje TC, Parsons JA, Sorenson RL 1994 Regulation of islet beta-cell proliferation by prolactin in rat islets. Diabetes 43:263-273 51. Zechner R, Strauss JG, Haemmerle G, Lass A, Zimmermann R 2005 Lipolysis: pathway under construction. Curr Opin Lipidol 16:333-340 52. Yeaman SJ 2004 Hormone-sensitive lipase--new roles for an old enzyme. Biochem J 379:11-22 53. Holm C 2003 Molecular mechanisms regulating hormone-sensitive lipase and lipolysis. Biochem Soc Trans 31:1120-1124

145 Figure Legends

Fig 1. Left panel: Determination of serum PRL levels in wild-type (+/+), heterozygous (+/-), and PRL knockout (-/-) mice using the Nb2 assay. Nb2 cells, rat lymphocytes which proliferate in the presence of PRL in a dose-dependent manner, were incubated with increasing concentrations of mPRL or mouse serum (0.5, 1, and 2 μl) for three days. Optical density was determined by the MTT assay. Inset. Serum PRL levels were calculated from the standard curve. Each value is a mean±SEM, n = 8 mice per group. M- males; F- females; ND- non detectable. Right panel: PRLR gene expression in liver and peri-uterine adipose tissue of female mice, as determined by real time RT-PCR. Values were calculated as relative gene expression vs that in wild-type mice. Each value is a mean±SEM, n = 8 mice per group. Similar results were obtained in males.

Fig 2. Growth curves of female (A) and male (B) wild-type (+/+), heterozygous (+/-), and PRL knockout (-/-) mice maintained on standard chow, low fat, or high fat diets. Each value is a mean±SEM, n = 8-10 for females and 20-30 for males.

Fig 3. Body composition of female and male wild-type (+/+), heterozygous (+/-), and PRL knockout (-/-) mice maintained on high fat or low fat diet for 12-14 weeks. Left and right panels display percent body fat and lean tissue, respectively, as determined by EchoMRI. Each value is a mean±SEM, n = 8-10 for females and 20-30 for males.

Fig 4. Serum lipids profile of male wild-type (+/+), heterozygous (+/-), and PRL knockout (-/-) mice maintained on high fat (HF), standard chow (SC) or low fat (LF) diet for 12-14 weeks. Blood was collected after an overnight fast by cardiac puncture and analyzed for phospholipids, triglycerides, cholesterol, and free fatty acids (FFA). Each value is a mean±SEM, n = 8 mice per group.

Fig 5. Serum leptin and adiponectin concentrations in male wild-type (+/+), heterozygous (+/- ), and PRL knockout (-/-) mice maintained on high fat (HF) or low fat (LF) diet for 12- 14 weeks. Blood was collected after an overnight fast by cardiac puncture and analyzed for leptin and adiponectin by the respective mouse-specific ELISA kits. Each value is a mean±SEM, n = 8 mice per group. * p < 0.05 vs. +/+ or +/- mice

Fig 6. Glucose tolerance determination in wild-type (+/+), heterozygous (+/-), and PRL knockout (-/-) male mice at 4, 8, or 12 weeks of age. Following an overnight fast, glucose levels were measured in blood collected from the tail vein before and after an ip injection of 10% D-(+)-glucose (10 µl/g body weight). Each value is a mean±SEM, n = 8-13 mice per group. At 4 weeks of age, the area under the curve was significantly (p<0.05) higher in -/- than +/+ mice.

Fig 7. Effects of PRL on basal- and isoproterenol-stimulated lipolysis in adipose tissue explants from mice, rats and humans. Epididymal (mice and rats) and subcutaneous abdominal (non-obese women) explants were incubated with PRL (0, 1, 5, and 25, ng/ml) for 24 hrs. Lipolysis was determined as described in Materials and Methods. Data are expressed as nmol glycerol/mg tissue/2 hours. Each value is a mean±SEM of 5 replicates. * p < 0.05 vs.

146 control explants. Each experiment was repeated 2-3 times with similar results. A similar effect of PRL on lipolysis was seen with subcutaneous abdominal explants obtained from non-obese men.

147 0.75 2 mPRL Adipose +/+ Liver -/- 0.50

Serum PRL (ng/ml) 1 -/- +/- +/+ 0.25 M ND 9.5±4.7 12.5±3.5

Optical Density F ND 28.0±9.3 23.1±7.6 Fold induction vs. +/+ vs. induction Fold .5 1 2 μl 0.00 0 /+ /- /- /+ /- - 0 1530456075 + + - + + -/ PRL (pg/well) Genotype

Figure 1

148 A Standard Chow Low Fat High Fat 30 30 30 +/+ +/- 25 25 -/- 25

20 20 20

15 15 15 Body weight (g)

10 10 10 4 8 12 16 4 8 12 16 20 4 8 12 16 20

B 40 40 40

30 30 30

20 20 20 Body weight (g)

10 10 10 4 8 121620 4 8 1216204 8 12 16 20 Age (weeks)

Figure 2

149 Females

100 20 Low Fat 80 High Fat 15 60 10 40

% Body Fat 5 20 % Lean Tissue

0 0 +/+ +/- -/- +/++/--/- Males 25 100 Low Fat 20 High Fat 80

15 60

10 40

% Body Fat 5 20 % Lean Tissue

0 0 +/+ +/- -/- +/+ +/- -/-

Figure 3

150 Phospholipids Triglycerides

500 HF SC LF 200

400 150 300 100

mg/dl 200 mg/dl 50 100

0 0

------+ - /- + - /- + /- /- /+ / -/ /+ / -/ /+ / -/ /+ / - /+ / - / + - + + + + + + + + + + + + 250 Cholesterol 2.00 FFA

200 1.50 150 1.00

mg/dl 100 mEq/dl 0.50 50

0 0.00

------/+ / -/ /+ / -/ /+ / -/ /+ / -/ /+ / -/ /+ / -/ + + + + + + + + + + + +

Figure 4

151 Leptin Adiponectin 25 15 HF 20 LF 10 15 ng/ml 10 µg/ml * 5 5

0 0 ------+ /- /- /+ / -/ /+ / -/ /+ / -/ / - + + + + + + + +

Figure 5

152 4 Weeks 8 Weeks 12 Weeks 500 500 500 +/+ +/ - 400 -/- 400 400

300 300 300

200

Glucose (mg/dl) 200 200

100 100 100 0 306090120 0 3060901200 306090120 Time (min)

Figure 6

153 Mouse (-/-) Mouse (+/+) 90 90 Basal Isoproterenol 70 70

50 50

30 30

Glycerol (nmol)/mg/2 hr 10 10 01525 01525 Rat Human 90 0.40

70 * * * 0.30 50 * * 0.20 30 * * * * *

Glycerol (nmol)/mg/2 hr 10 0.10 01525 01525 PRL (ng/ml) PRL (ng/ml)

Figure 7

154

Chapter 4. Expression of Selected Genes in PRL-Deficient Mice

155 Introduction

Proper regulation of metabolic homeostasis is crucial for survival, and involves a number of pathways whose components are expressed and activated depending upon the organism's energy needs and physiological activity. Adipose tissue and the liver play central roles in this process, producing enzymes and hormones that ensure proper utilization and mobilization of carbohydrates and lipids during widely varying physiological states, including the fed state and during fasting.

Leptin and fasting-induced adipocyte factor (FIAF) are adipokines whose serum levels decrease and increase in response to fasting, respectively. Reflective of depleting energy stores, low levels of leptin leads to increased food intake and decreased energy expenditure (1). Despite several studies, the relationship between PRL and leptin production remains unclear (2-4). FIAF, which is secreted from both liver and adipose tissue, inhibits TG uptake by blocking lipoprotein and chylomicron binding sites on LPL (5). FIAF exists as at least three isoforms, a native 50 kDa protein, and two short isoforms resulting from cleavage of the native protein. These isoforms, deemed S1 and S2, are both N-terminal fragments of ~35 kDa and are differentially expressed in several tissues, including adipose tissue and liver (6;7). It is not entirely clear which FIAF isoform is responsible for its physiological actions, however one study has reported that levels of the N-terminal isoforms correlate well with plasma TG (8). Although PRL has long been implicated in lipid metabolism, no studies have investigated its effects on FIAF. On the other hand, PRL has been reported to inhibit secretion of certain adipokines, including adiponectin, which is involved in obesity-related insulin resistance (9;10).

Lipoprotein lipase (LPL) and hormone sensitive lipase (HSL) are the major enzymes controlling lipid influx and efflux (lipolysis), respectively. Limited reports suggest that PRL regulates both enzymes. One study has shown that PRL inhibits LPL activity in humans, while there are conflicting reports on whether PRL is lipolytic (11-13). PRLR signaling activates several pathways that mediate gene transcription, and PRL has been shown to affect the expression of several metabolic factors, including fatty acid synthase and PPARγ (14;15). No studies have investigated the effects of PRL on gene expression of most of the aforementioned metabolic factors. To address this issue, we measured the expression of selected genes in adipose tissue and liver of PRL-deficient and wild-type mice under fasted and fed conditions. We then analyzed the protein levels of any genes that differed between the genotypes.

Materials and methods

Animals

PRL-knockout, heterozygous, and wild-type mice were bred and housed as described in chapter 3, with standard chow and water available ad libitum unless otherwise specified. Animals were either allowed free access to food or were fasted for 24 hrs before sacrifice by CO2 inhalation and cervical dislocation between 0900 and 1100 h. Blood was collected by cardiac puncture and centrifuged at 5000 x g for 10 min at 4 C. Serum was kept at –80 C for later analysis. Liver and adipose tissues (epididymal or periuterine) were removed and frozen for analysis by RT-PCR.

156 Real Time PCR

Total RNA was extracted, reverse transcribed and analyzed by real time PCR as described in chapter 3. Product purity was confirmed by DNA melting curve analysis. β-Actin was used as a reference gene. Fold changes in gene expression were calculated from the cycle threshold measurements, using the method of Pfaffl et al. (16).

Western blotting

For FIAF determination, 0.5 μl of serum was subjected to electrophoresis on 12% SDS-PAGE gels and the separated proteins were transferred onto nitrocellulose membranes. Membranes were washed 2X with PBST and then incubated in Odyssey blocking buffer (1:1 in PBST) for 1 hour at room temperature. Membranes were incubated overnight at 4°C with primary FIAF antibody (1:1000), followed by three 15 minute washes with PBST. Incubation with a fluorescent secondary antibody at 1:10000 (Alexa Fluor 700, Molecular Probes, Invitrogen, Carlsbad CA) was done at room temperature for 45 minutes. Proteins were visualized and quantified using an Odyssey Infrared Imaging System (Li-Cor Biosciences, Lincoln NE).

Data analysis

For real time PCR, gene expression was analyzed using REST (relative expression software tool V1.9.9) (16). For quantitative western blotting, statistical differences were determined by one- way ANOVA followed by post hoc analysis comparing least significant differences. P values <0.05 were considered significant.

Results

PRL-deficiency has no effect on selected gene expression in fed mice

To determine the effects of PRL-deficiency on gene expression, mRNA from adipose tissue and liver was quantified using RealTime PCR. β-actin was used as a positive control. The primer pairs used in these studies are presented in Table 1, and gene expression data are shown in Table 2. Each value is presented as relative expression vs. fed wild-type animals. As seen in the table, PRL deficiency had no affect on any of the genes analyzed under fed conditions.

Fasting increases FIAF expression in the liver and adipose tissue of wild-type male mice, but not PRL-deficient mice

As shown in Fig 1. left panel, fasting dramatically increased FIAF expression in the liver of wild-type male mice, but not in male PRL-knockouts. FIAF expression was also significantly elevated by fasting in adipose tissue, though not as marked as in the liver of wild-type males (right panel). This effect was not observed in adipose tissue of fasted male PRL-knockouts. Fasting increased FIAF expression in the liver, but not adipose tissue of females regardless of genotype (not shown). As shown in Fig 2, fasting significantly decreased the expression of leptin (left panel) and LPL (right panel) in adipose tissue of males regardless of genotype. Similar results were seen in females (data not shown).

157

Elevated concentration of two FIAF isoforms in serum of PRL-deficient male mice

We next questioned whether circulating FIAF levels differ between wild-type and PRL-deficient mice under fed and fasted conditions. To address this issue, we used western blotting and compared serum FIAF concentrations. As shown in Fig 3. Both the S1 (upper panel) and S2 (lower panel) isoforms were present at higher levels in PRL-deficient mice when compared to wild-types. As shown in Fig 4, following scanning and expression as fluorescent units, both isoforms were present at ~50% higher levels than in wild-type mice. Fasting had no effect on the expression of either isoform within the same genotype. No differences were observed in the concentration of either isoform in females regardless of genotype (not shown).

Discussion

Here we show that show that 1) fasting increases FIAF expression in the liver and adipose tissue of wild-type, but not PRL-deficient male mice, and 2) the concentration of two short FIAF isoforms is elevated in the serum of PRL-deficient male mice when compared to wild-type mice. This effect was not seen in females.

Because the metabolic status of an organism depends on its nutritional as well as hormonal status, we analyzed tissue and serum from fed and fasted animals. In the fed state, PRL deficiency had no effect on any genes measured in either tissue. These data indicate that when sufficient energy is available, PRL-deficiency has no apparent effect on some metabolic-related genes expressed in adipose tissue or liver. Previous in vitro studies claiming that PRL regulates HSL used extremely high doses of PRL (5μg/ml) (12)

Both leptin and LPL decrease as energy levels drop in order to decrease energy expenditure and lipid storage in adipose tissue. Indeed, fasting caused down-regulation of leptin, LPL and several other genes in adipose tissue of our mice. These observations are important, since they validate our experimental model. However, PRL deficiency had no effect on the expression of these genes. While PRL has been reported to inhibit LPL activity in adipose tissue explants in vitro, those studies used human tissue; and as we have shown in chapter 3, the metabolic actions of PRL often vary among species (11). As also discussed in the previous chapter, PRLR-deficiency reportedly results in significantly decreased serum leptin levels; however, this was attributed to decreased adipose tissue mass, and likely not due to changes in gene expression (17). However, more recent studies have reported unaltered circulating leptin levels in PRLR-deficient mice (18).

As expected, fasting significantly increased FIAF expression in adipose tissue and liver of male mice, as well as in the liver of females. Notably, fasting failed to induce FIAF mRNA levels in male PRL-deficient mice, suggesting that PRL is necessary for the fasting induced increase of FIAF by a yet to be identified regulatory mechanism. Additionally, we are uncertain why the fasting response is not seen in adipose tissue of females. Previous studies that investigated the effects of fasting on FIAF expression have yet to indicate which gender this response occurs in.

158 Our analysis of serum FIAF levels yielded interesting results. Similar to another report, we did not observe a response to fasting in the S1 and S2 isoforms (19). While the short isoforms are cleavage products of the native 50 kDa isoform, they are differentially expressed, with adipose tissue producing S1 and S2, and liver producing S1 and native isoforms. Thus, we speculate that PRL may act on either WAT, or both WAT and liver to alter the expression of these isoforms.

It is unclear which FIAF isoform is responsible for its physiological actions. Only one study has addressed this issue, reporting that FIAF was cleaved in vivo, and levels of the N-terminal isoform correlated with plasma TG. This suggests that the N-terminal fragment mediates biological activity and raises the notion that proteolytic processing of FIAF is a mechanism by which its activity is regulated. This idea is intriguing and warrants further investigation.

Since the antibody used in our studies detects N-terminal fragments of FIAF, it is possible that alterations in FIAF are representative of its biological activity. Given the ability of FIAF to elevate plasma TG levels by inhibiting LPL activity, one might expect higher TG levels in PRL- deficient mice. However, we did not observe any changes in plasma TG between PRL-deficient and wild-type mice, and it is possible that a ~50% increase in FIAF levels is insufficient to elicit a response. Indeed, all studies to date on FIAF have used overexpression models, making it unclear what concentrations of FIAF are necessary to exert its biological effects.

159 References

1. Woods SC, Benoit SC, Clegg DJ, Seeley RJ 2004 Clinical endocrinology and metabolism. Regulation of energy homeostasis by peripheral signals. Best Pract Res Clin Endocrinol Metab 18:497-515 2. Gualillo O, Lago F, Garcia M, Menendez C, Senaris R, Casanueva FF, Dieguez C 1999 Prolactin stimulates leptin secretion by rat white adipose tissue. Endocrinology 140:5149- 5153 3. Ling C, Billig H 2001 PRL receptor-mediated effects in female mouse adipocytes: PRL induces suppressors of cytokine signaling expression and suppresses insulin-induced leptin production in adipocytes in vitro. Endocrinology 142:4880-4890 4. Viengchareun S, Bouzinba-Segard H, Laigneau JP, Zennaro MC, Kelly PA, Bado A, Lombes M, Binart N 2004 Prolactin potentiates insulin-stimulated leptin expression and release from differentiated brown adipocytes. J Mol Endocrinol 33:679-691 5. Kersten S 2005 Regulation of lipid metabolism via angiopoietin-like proteins. Biochemal Society Transections 33:1059-1062 6. Kersten S, Mandard S, Tan NS, Escher P, Metzger D, Chambon P, Gonzalez FJ, Desvergne B, Wahli W 2000 Characterization of the fasting-induced adipose factor FIAF, a novel peroxisome proliferator-activated receptor target gene. Journal of Biological Chemistry 275:28488-28493 7. Kim I, Kim HG, Kim H, Kim HH, Park SK, Uhm CS, Lee ZH, Koh GY 2000 Hepatic expression, synthesis and secretion of a novel fibrinogen/angiopoietin-related protein that prevents endothelial-cell apoptosis. Biochem J 346 Pt 3:603-610 8. Ge H, Yang G, Yu X, Pourbahrami T, Li C 2004 Oligomerization state-dependent hyperlipidemic effect of angiopoietin-like protein 4. J Lipid Res 45:2071-2079 9. Combs TP, Berg AH, Rajala MW, Klebanov S, Iyengar P, Jimenez-Chillaron JC, Patti ME, Klein SL, Weinstein RS, Scherer PE 2003 Sexual differentiation, pregnancy, calorie restriction, and aging affect the adipocyte-specific secretory protein adiponectin. Diabetes 52:268-276 10. Nilsson L, Binart N, Bohlooly Y, Bramnert M, Egecioglu E, Kindblom J, Kelly PA, Kopchick JJ, Ormandy CJ, Ling C, Billig H 2005 Prolactin and growth hormone regulate adiponectin secretion and receptor expression in adipose tissue. Biochem Biophys Res Commun 331:1120-1126 11. Ling C, Svensson L, Oden B, Weijdegard B, Eden B, Eden S, Billig H 2003 Identification of functional prolactin (PRL) receptor gene expression: PRL inhibits lipoprotein lipase activity in human white adipose tissue. J Clin Endocrinol Metab 88:1804-1808 12. Fielder PJ, Talamantes F 1987 The lipolytic effects of mouse placental lactogen II, mouse prolactin, and mouse growth hormone on adipose tissue from virgin and pregnant mice. Endocrinology 121:493-497 13. Fortun-Lamothe L, Langin D, Lafontan M 1996 Influence of prolactin on in vivo and in vitro lipolysis in rabbits. Comp Biochem Physiol C Pharmacol Toxicol Endocrinol 115:141-147 14. Hogan JC, Stephens JM 2005 The regulation of fatty acid synthase by STAT5A. Diabetes 54:1968-1975

160 15. Stewart WC, Baugh JE, Jr., Floyd ZE, Stephens JM 2004 STAT 5 activators can replace the requirement of FBS in the adipogenesis of 3T3-L1 cells. Biochem Biophys Res Commun 324:355-359 16. Pfaffl MW, Horgan GW, Dempfle L 2002 Relative expression software tool (REST) for group-wise comparison and statistical analysis of relative expression results in real-time PCR. Nucleic Acids Res 30:e36 17. Freemark M, Fleenor D, Driscoll P, Binart N, Kelly P 2001 Body weight and fat deposition in prolactin receptor-deficient mice. Endocrinology 142:532-537 18. Fleenor D, Oden J, Kelly PA, Mohan S, Alliouachene S, Pende M, Wentz S, Kerr J, Freemark M 2005 Roles of the lactogens and somatogens in perinatal and postnatal metabolism and growth: studies of a novel mouse model combining lactogen resistance and growth hormone deficiency. Endocrinology 146:103-112 19. Mandard S, Zandbergen F, Tan NS, Escher P, Patsouris D, Koenig W, Kleemann R, Bakker A, Veenman F, Wahli W, Muller M, Kersten S 2004 The direct peroxisome proliferator-activated receptor target fasting-induced adipose factor (FIAF/PGAR/ANGPTL4) is present in blood plasma as a truncated protein that is increased by fenofibrate treatment. Journal of Biological Chemistry 279:34411-34420

161 Gene Accession # Sense primer Antisense primer ACTGCTCTGGCTCCT AGTCCGCCTAGAAGC β-Actin NM_007393 AGCAC ACTTG CACAGTAAATGCCAC GGCAACCATTTTACC PRLR NM_011169 GAACG CACAG TGACACCAAAACCCT TCATTGGCTATCTGC Leptin NM_008493 CATCA AGCAC CAGCTCCTGTCATTC AGCTAGCTCCTGCTT Adiponectin NM_009605 CAACA TGGTC TTTTCTGGGACTGAG TCATGAGCAGTTCTC LPL NM_008509 GATGG CGATG TGGATGTGCACTTCT CCAAGGGAGGTGAG HSL MMU08188 GGAAA ATGGTA CTTCAGAGCCAGATT AAATGGTGGAGATCC FIAF NM_020581 GACCT CAGAG GCTGTTATGGGTGAA GGCTTGATGTCAAAG PPARγ NM_011146 ACTCTGGGA GAATGCGAG TCATTTCCCCTCCTTT CAAGACTGCTGTGCC Resistin NM_022984 TCCT TTCTG CACAGAAGGTGATTG TTTCCTTCCCAACCA Glut4 AB008453 AACAG TTGAG GCTACACCTGAAGAC GGAGACCAATTGCAC PRL NM_011164 AAGGA CCAA

Table 1. PCR primers for selected mouse genes. Abbreviations: PRLR, prolactin- receptor; LPL, lipoprotein liase; HSL, hormone sensitive lipase; FIAF, fasting-induced adipocyte factor; PPARγ, peroxisome proliferator-activated receptor; Glut4, glucose transporter 4; PRL, prolactin

162 Fed Fasted Male +/+ -/- ±SE M +/+ ±SEM -/- ±SEM Liver PPARγ 1 1.31 0.65 1.47 1.24 1.76 1.12 LPL 1 0.61 0.38 1.04 0.32 1.19 0.47 Adipose tissue PPARγ 1 1.57 0.70 0.78 0.15 0.90 0.05 HSL 1 0.88 0.20 0.88 0.33 1.57 0.78 Adiponectin 1 0.88 0.22 0.68 0.13 0.67 0.32 Resistin 1 0.96 0.12 0.45 0.34 0.76 0.23

Female +/+ -/- ±SE M +/+ ±SEM -/- ±SEM Liver FIAF 1 1.23 0.47 5.23 2.73 8.43 3.39 PPARγ 1 1.49 0.88 1.94 1.07 0.64 0.31 LPL 1 1.07 0.10 2.23 0.55 2.92 0.28 Adipose tissue FIAF 1 1.49 0.87 1.94 1.07 0.98 0.28 PPARγ 1 0.67 0.26 2.09 1.45 1.50 0.12 LPL 1 0.85 0.35 0.52 0.20 0.51 0.13 HSL 1 0.66 0.21 1.92 1.14 0.98 0.21 Adiponectin 1 0.68 0.25 0.65 0.11 0.78 0.31 Resistin 1 1.33 0.23 0.45 0.12 0.55 0.22 Leptin 1 0.74 0.08 0.30 0.05 0.29 0.17

Table 2. Expression of selected genes in the adipose tissue and liver of wild- type (+/+) and PRL-deficient (-/-) mice under fed or 24 hour fasted conditions. Data are expressed as relative expression vs fed wild-type ±SEM. Value are representative of at least 4 animals.

163 Liver Adipose 20 20 Fed 16 16 Fasted

12 12

8 8

4 4

Relative expression vs fed +/+ * * 0 0 +/+ +/+ -/- -/- +/+ +/+ -/- -/-

Fig 1. Effects of PRL-deficiency on FIAF gene expression, as determined by real time PCR. mRNA was measured in the liver (left panel) and adipose tissue (right panel) of wild-type (+/+) and PRL-deficient male mice under fed conditions or following a 24 hour fast. Values are the mean of at least 4 animals per group and expressed as relative expression vs. fed wild-type mice. *p<0.05 vs. fasted +/+ mice.

164 Leptin LPL 1.50 1.50 Fed

Fasted

1.00 1.00

* 0.50 0.50 * * * Relative expression vs fed +/+

0.00 0.00 +/+ +/+ -/- -/- +/+ +/+ -/- -/-

Fig 2. Effects of PRL-deficiency on gene expression in adipose tissue of male mice, as determined by real time PCR. Leptin (left panel) and lipoprotein lipase (LPL) (right panel) mRNA levels were measured in the adipose tissue of wild-type (+/+) and PRL- deficient male mice under fed conditions or following a 24 hour fast. Values are the mean of at least 4 animals per group and expressed as relative expression vs. fed wild-type mice. *p<0.05 vs. fed +/+ mice.

165 MW Wild-type PRL-deficient 40kDa S1 25kDa S2

Fasted - + - + - + - + - + - + - +

Fig 3. Effect of PRL deficiency on FIAF serum levels in male mice, as assessed by western blotting. Both panels are different exposures of the same blot. Both the S1 (upper panel) and S2 (lower panel) isoforms are present at higher levels in PRL- deficient mice as compared to wild-types. Fasting had no effect on the concentration of either isoform.

166 S1 S2 1.50 20 Fed Fasted * * * * 15 1.00

10

Arbitrary units 0.50 5

0.00 0 +/+ -/- +/+ -/- +/+ -/- +/+ -/-

Fig 4. Effect of PRL deficiency on FIAF serum levels in male mice, as assessed by quantitative western blotting. Both the S1 (left panel) and S2 (right panel) isoforms are present at higher levels in PRL-deficient mice (-/-) as compared to wild-types (+/+). Fasting had no effect on the concentration of either the S1 or S2 isoform within the same genotype. Band intensity was determined using Odyssey imaging software, and data are expressed as arbitraty units. *p<0.05 vs +/+ mice

167

Chapter 5. The Role of PRL in Preadipocyte Proliferation and Lipid Accumulation

168 Introduction

White adipose tissue is unique in its potential for large volume changes, with fat mass increasing up to 4-fold in morbidly obese humans and reaching 60% to 70% of total body weight (1;2). WAT expansion results from increases in adipocyte size, caused by lipogenesis and lipid uptake, as well as increased cell number, a result of preadipocyte proliferation and differentiation. There are multiple factors contributing to the growth of adipose tissue, including, but not limited to dietary and hormonal factors. Both adipocyte hyperplasia and hypertrophy have both been observed in various rodent models of obesity, including rats fed a high-fat or high-carbohydrate diet (3;4).

There is considerable evidence of hormonal regulation of adipocyte growth and development. GH-receptor knockout mice exhibit major decreases in the number of parametrial adipocytes, while transgenic inactivation of GH in older normal mice results in an obese phenotype (5). When we performed the set of experiments discussed in this section, little data were available on the effects of PRL on adipose tissue growth. The lone report using PRLR-deficient mice suggested that PRL was a positive regulator of adipose tissue mass, based on the observation that abdominal fat content was reportedly as much as 50% lower than that in wild-type mice. A more detailed, but unreported analysis of these animals revealed that the decrease in adipose tissue mass was at least in part a result of decreased adipocyte number, thereby implicating PRL in adipocyte proliferation and/or differentiation (6). Murine 3T3-L1 preadipocytes are commonly used as a model for studying adipogenesis in vitro. In order to proliferate, these cells require the presence of FBS, which contains lactogenic hormones. PRL acts as mitogen in numerous cell types, including prostate and mammary epithelial cells. However, the role of PRL in preadipocyte growth has not yet been investigated.

Upon exposure to adipogenic media containing insulin, IBMX, and dexamethasone, 3T3-L1 cells differentiate from fibroblast-like preadipocytes, into mature, lipid-filled adipocytes. During this process, there exists coordinated induction of several adipogenic transcription factors including C/EBPs and PPARγ. At the time that our studies began, there were several lines of evidence suggesting that Stat proteins, and therefore PRL, play a role in adipogenesis. For example, it has been noted that the presence of FBS, which contains lactogenic hormones, was required to achieve efficient differentiation of 3T3-L1 cells (7). When differentiated in CSS, which is devoid of lactogens, these cells fail to fully differentiate. Because IBMX and dexamethasone are direct inducers for C/EBPß and -δ, respectively, it has therefore been hypothesized that factors activated by FBS play a role in adipogenesis. It has also been reported that expression and activation of Stat 5a and 5b are robustly upregulated during differentiation (8). Moreover, ectopic expression of Stat5a during early adipogenesis in NIH-3T3 cells upregulates PPAR-γ and increases triglyceride storage (7). Finally, expression of a dominant negative form of Stat 5a attenuates lipid accumulation in NIH-3T3 cells, as well as PPARγ expression in mice.

Both preadipocytes and mature adipocytes express several cytokine receptors, including the PRLR, which was recently detected in adipose tissue (9). Moreover, PRLR expression reportedly increases in differentiating bone marrow stromal cells and T37i brown adipocytes (10;11). This further supports the notion that PRL may play an effect in adipogenesis. Therefore, we hypothesized that PRL was a positive stimulator of preadipocyte proliferation and differentiation

169 in 3T3-L1 cells, which express higher levels of PRLR, and require fewer adipogenic cues than NIH-3T3 cells. Our goals were to: 1) demonstrate the expression of the PRLR in non- differentiated and differentiating cells, 2) determine the effects of PRL on 3T3-L1 preadipocytes proliferation under serum-free and lactogen deprived conditions, and 3) determine the effects of PRL on lipid accumulation and morphological changes during differentiation.

Materials and methods

Reagents

3T3-L1 cells were obtained from Dr. Philip Scherer, Albert Einstein College of Medicine. Trypsin, as well as high- and low-glucose Dulbecco’s modified Eagle medium (DMEM) were purchased from Gibco (Carlsbad, CA). Fetal bovine serum and Charcoal stripped serum (CSS) were purchased from JRH biosciences (Lenexa, KA) and Hyclone (Logan, UT), respectively. Dexamethasone, 1-methyl-3-isobutylxanthine (IBMX), bovine insulin, and sodium dodecyl sulfate (SDS) were obtained from Sigma-Aldrich (St. Louis, MO). Recombinant ovine PRL was a gift from Dr. Arieh Gertler, The Hebrew University of Jerusalem.

Cell culture

3T3-L1 preadipocytes were maintained in low-glucose Dulbecco’s modified Eagle medium (DMEM) (Gibco, Carlsbad, CA) containing 5% FBS (JRH biosciences, Lenexa, KA). Cells were only used at low passage, and split every two days to prevent them from reaching confluence.

Cell viability assays

For proliferation experiments, preadipocytes were seeded in 96-well plates (1000 or 4000 cells/well) in DMEM containing 1% CSS. After overnight incubation, cells were treated with PRL in either 1% CSS or SFM. 5% FBS served as a positive control in all experiments. After 1, 2, or 4 days, cell viability was determined using either MTT, Calcein AM, or Resazurin.

For the MTT assay, cell media was removed and 20 μl of (3-[4,5-dimethylthiazole-2-yl]-2,5- diphenyltetrazolium bromide (Sigma) MTT was added to each well. After 2 hr incubation at 37°C in a 5% CO2 environment, the MTT was removed and 100 μl of dimethyl sulfoxide (DMSO) was added to each well. Optical density at 570nm was then read on a microplate reader (Biotek Instruments Inc., Winooski VT).

For Calcein AM assay, media was removed and Calcein AM was added at a final concentration of 1µg/ml. Plates are wrapped in foil and incubated at 37°. Reading occurred at a few time points between 20 min and an hour (30 min or longer probably best) using the fluorimeter (485 nm excitation and 515 nm emission).

For the resazurin assay, media was removed and resazurin (made in PBS) was added at a final concentration of 12.5 µg/ml. After a 2 hr incubation at 37°, fluorescence is read at 530 nm excitation and 590 nm emission, using Spectramax Gemini XPS fluorimeter (Molecular Devices, Sunnyvale, CA).

170

3T3-L1 differentiation

For differentiation experiments, preadipocytes were plated at 11,000 cells/cm2 in 96 well plates or chamber slides, and grown to confluence. Two days later, cells were induced to differentiate by changing the medium to high-glucose DMEM with 0.5 mM 3-isobutyl-methylxanthine (IBMX), 0.5 µM dexamethasone (DEX), 10μg/ml insulin, and 5% CSS +/- PRL. 5% FBS was used as a positive control. After 48 h, this medium was replaced with DMEM containing the same treatment, but without IBMX, DEX, and insulin. Cells differentiated in CSS were supplemented with biotin and pantothenate, factors that are necessary for lipogenesis but removed during charcoal/dextran treatment. Media was changed every 2 days until photos were taken or TG measured.

Oil red O staining and measurement of lipid accumulation

Adipocytes were fixed in 4% paraformaldehyde for 30 minutes, rinsed with 10 mM PB, then 70% EtOH. Cells were then stained with Oil Red O (5mg/200ml isopropanol, diluted 6:4 in ddH20, and then filtered) for 60 minutes. To measure lipid content, cells were stained with Oil Red O and then rinsed with ddH2O. Lipids were extracted with 100ul of isopropanol and optical density was measure using a microplate reader. For cells that were photographed, chamber slides were rinsed with 70% EtOH after Oil Red O, and nuclei were stained with hematoxylin for 1-2 minutes. Subsequently, cells were rinsed with ddH20 and then with warm tap water before being coverslipped. Slides were photographed using a SPOT-2 digital CCD camera mounted on a Nikon Microphot microscope (Nikon Corp., Melville NY).

Real Time PCR

Total RNA was extracted, reverse transcribed and analyzed by real time PCR as described in chapter 3. Intron-spanning primers for GLUT4 and β-Actin were used. Cycle parameters were 96 C for 6 min for polymerase activation, followed by 40 cycles of 95 C for 15 sec, 57 C for 15 sec, and 72 C for 25 sec, and finally an optical read stage at 83.5 C for 6 sec. Product purity was confirmed by DNA melting curve analysis. β-Actin was used as a reference gene. Fold changes in gene expression were calculated from the cycle threshold measurements, using the method of Pfaffl et al. (12).

Data analysis

For real time PCR, gene expression was analyzed using REST (relative expression software tool V1.9.9) (12). For quantitative western blotting, statistical differences were determined by one- way ANOVA followed by post hoc analysis comparing least significant differences. P values <0.05 were considered significant.

Results

3T3-L1 preadipocytes maintained in 5% CSS grow slower than in 5% FBS

171 The MTT, resazurin, and Calcein AM assays were used to determine cell viability. As shown in Fig 1, cell number (from 250-8000 cells/well) showed a linear relationship with optical density, as determined using the MTT assay (A), as well as with relative fluorescent units as determined by resazurin (B) and calcein AM (C) assays. To demonstrate the difference in growth rate, cells (2000/well) were incubated in the presence of 5% CSS or 5% FBS for 4 days, and cell number was assessed every 24 hours using the three viability assays. As shown in Fig 2, cells maintained in 5% FBS grew at a significantly higher rate than those incubated in 5% CSS. The doubling time of cells grown in FBS was approximately 48 hours, while in CSS it was 96 hours, as determined by the (A) MTT, (B) rezasurin, and (C) calcein AM assays.

PRL has insignificant effects on 3T3-L1 preadipocyte growth

To determine the effects of PRL on preadipocyte proliferation, non-differentiated 3T3-L1 cells were plated at a low density and treated with increasing doses of PRL in either 1% CSS or SFM. FBS served as a positive control in all experiments. Cell viability was determined using the assays described above. As shown in Figs. 3 & 4, preadipocytes grown in SFM (left panel) exhibited significantly lower viability than those exposed to 1% CSS (right panel), while cells incubated in the presence of 5% FBS maintained a high level of viability in all experiments. 48 hr PRL treatment had no effect on preadipocytes incubated in the presence of SFM; however when treated in the presence of 1% CSS, PRL caused a dose-dependent, yet insignificant increase in cell viability. This effect was not observed in all experiments, and higher doses of PRL (250 ng/ml), did not increase this effect. In addition, treatment for 24, and 96 hours had no effect on viability when compared to control cells, as assessed by all assays (data not shown).

Effects of PRL on morphological changes and lipid accumulation during adipogenesis

We next questioned whether PRL plays a role in adipogenesis. To address this issue, we first demonstrated expression of PRLR in 3T3-L1 cells. Preadipocytes were differentiated in the presence of FBS, and expression was assessed by RT-PCR. As shown in Fig. 5, upper panel, PRLR mRNA levels were low but detectable on day 0 (lane 1), and significantly increased by day 2 (lane 2). PRLR expression continued to rise by day 4 (lane 3), and remained high throughout differentiation (lanes 4 and 5). PRL expression was also examined, but was not detected at any time (upper lane). Murine pituitaries were used as a positive control for both genes (lane 6). We also confirmed the presence of the PRLR in our experimental model of adipogenesis by realtime PCR, which is performed in the presence of CSS rather than FBS. As shown in Fig 5, lower panel, PRLR levels are low prior to differentiation, increase significantly by day 2, and remain relatively steady thereafter.

To assess the role of PRL in adipogenesis, 3T3-L1 cells were differentiated in the presence or absence of PRL for 8 days and lipid accumulation was measured every 48 hours. As shown in Fig. 6, preadipocytes differentiated in the presence of FBS accumulated TG slowly during the first four days, then significantly increased lipid storage during days 4-8. When differentiated in CSS, there was virtually no lipid accumulation during days 0-4, with only a slight increase thereafter. By day 8, these cells contain approximately as much as those treated with FBS accumulate by day 4. 25ng/ml PRL had had no affect on lipid storage during differentiation, with

172 cells accumulating the same amount of TG as those treated with CSS alone. Similar results were obtained when cells were differentiated in 125 and 250 ng/ml (not shown).

These results were confirmed by microscopic analysis of 3T3-L1 cells stained with Oil Red O and hematoxylin. As shown in Fig. 5, cells on day 0 were fibroblast-like in shape and contained no lipid droplets. Two days after the induction of differentiation, there is little change in cell morphology, with no differences between treatment groups (not shown). At day 4, cells differentiated in FBS are large and contain numerous lipid droplets, while those differentiated in CSS +/- PRL (25 ng/ml) are smaller, with little amounts of lipid stored. After 8 days, adipogenic media containing FBS induced the formation of large, round adipocytes that display an abundant amount of Oil Red O staining. Similarly, cells differentiated in CSS appeared round in shape, but were smaller and accumulated far fewer lipid droplets after 8 days. Again, PRL had no effect on either cell morphology or the amount of stored lipid.

Discussion

In this section, we are reporting that 1) PRL treatment had no significant effects on preadipocyte proliferation, 2) PRLR expression increases during 3T3-L1 differentiation, and 3) PRL does not affect lipid accumulation or expression of adipogenic markers during differentiation.

To validate our experimental design, we first demonstrated that cell number correlated with optical density as measured by the MTT assay, as well as relative fluorescent units as determined using resazurin and calcein AM. Based on these data we determined an optimal plating density for our studies, which falls in the linear range of the curve.

As precursors for mature adipocytes, preadipocytes regulate the capacity for growth within adipose tissue. Here we have shown that 3T3-L1 preadipocytes proliferate at a much slower rate in CSS when compared to FBS, as determined by all three assays. This observation lead us to hypothesize that PRL, which is removed during charcoal/dextran treatment, is a serum component necessary for cell growth. PRL is a potent mitogenic factor in several cell types, including prostate and mammary epithelial cells (13;14). PRL caused a small, but insignificant increase in cell viability when CSS was present, but not SFM, in our initial experiments. These data suggest that while CSS alone is sufficient to maintain the viability of preadipocytes, PRL can further induce cell growth. However, this dose-dependent effect was not further increased using higher doses of PRL (250 ng/ml). Additionally, subsequent studies showed that PRL treatment had no effect on preadipocyte growth, as determined by two additional cell viability assays.

While MTT is an indicator of mitochondrial succinate dehydrogenase activity, which reduces FAD to FADH2, rezasurin reflects cytoplasmic diaphorase activity, representative of NAD(P) reduction. Additionally, calcein AM is an indicator of esterase activity. Although these assays are used to measure cell viability, they each represent a unique biochemical event. Thus, it is possible that PRL could alter the levels of one metabolic parameter, succinate dehydrogenase in the case of our observations, but not affect activity of other enzymes, and subsequently cell growth. While this may be the case, the effects of PRL were minor; leading us to believe that preadipocyte proliferation requires the presence of additional serum component. This yet to be

173 identified factor(s) is likely removed during charcoal stripping and is crucial for normal cell growth. A candidate molecule could be one of the prostoglandins, which are significantly reduced by charcoal/dextran treatment, and have been shown to initiate proliferation in NIH-3T3 preadipocytes and cultured mouse fibroblasts (15-17).

It is also possible that the effects of PRL vary among preadipocyte cell types, with 3T3-L1 being a relatively PRL-unresponsive cell line. Indeed, GH has been reported to decrease proliferation in murine 3T3-F442A preadipocytes, but stimulate the growth of preadipocytes in primary cultures from rat and human adipose tissue (6). Moreover, in other tissues such as the prostate gland, PRL exhibits regional mitogenic effects (18). Because of their insensitivity it is possible that PRL at much higher doses than those used in these studies would affect 3T3-L1 cell proliferation or differentiation.

There are several lines of evidence that suggest PRL plays a role in adipogenesis. For instance, the PRLR is expressed in both mouse and human adipose tissue and is induced during differentiation of bone marrow stromal cells and T37i brown adipocytes (10;11). In agreeance with these reports, we observed increased levels of PRLR mRNA during 3T3-L1 adipogenesis. Expression was low but detectable prior to differentiation, but dramatically increased within 48 hours of exposure to adipogenic media.

Growth factors such as insulin, IGF-1, epidermal growth factor (EGF), and platelet-derived growth factor promote adipogenesis in 3T3-L1 cells, with FBS reported to be a potent stimulus (19-21). Indeed, we observed that induction of differentiation in CSS causes few morphological changes and only a small amount of lipid accumulation. That these cells did not differentiate well in the absence of FBS suggests that a serum component is necessary for efficient conversion to adipocytes. PRL is a candidate factor in this process, since expression and activation of Stat 5 proteins correlates with adipogenesis and lipid accumulation (8). In our experiments, addition of PRL to adipogenic media had no effect on either lipid storage or cell morphology. This does not agree with some reports that have been published since the time we performed our studies.

Notably, after our studies were completed, one group reported that Stat5 activators, particularly PRL and GH, can replace the requirement of FBS during 3T3-L1 differentiation (22). However, these studies used extremely high doses of PRL (>0.5 μg/ml) and GH (>1 μg/ml). Furthermore, while lipid staining was used as a measure of adipogenesis induced by GH, differentiation in PRL-treated cells was assessed only by the expression of PPARγ, an early adipogenic marker. In fact, these authors state that “some other factor present in calf serum is needed for PRL and GH to induce maximal differentiation”. Therefore, PRL still has not been demonstrated to induce lipid accumulation or morphological characteristics acquired during mid- to late-adipogenesis.

Another study reported that the addition of PRL to normal adipogenic media enhanced expression of PPARγ, but had no effect on terminal differentiation marker gene expression or morphological changes in NIH-3T3 cells (23). Only when cells over-expressing PRLR and treated with extremely high doses of PRL (1 μg/ml), as well as the PPARγ ligand troglitazone, did a modest number (13% above control) of cells form lipid droplets.

174 Lipid accumulation is a hallmark characteristic of adipogenesis, reflecting the combined rates of lipid uptake via LPL, de novo lipogenesis by FAS, and lipolysis by HSL. LPL is upregulated during early differentiation, while the importance of FAS in adipocyte function is underscored by the inhibition of adipogenesis by C75, an allosteric inhibitor of FAS (24). Interestingly, PRL downregulates FAS mRNA and protein levels in 3T3-L1 adipocytes via a Stat5 dependent mechanism, and inhibits LPL activity in human adipose tissue (25;26). Since PRL is reported to be a pro-adipogenic factor, it is paradoxical that PRL has also been shown to down-regulate expression and/or activity of lipogenic machinery. In fact, these data could lead to the assumption that PRL exhibits anti-adipogenic activity.

Taken together, it appears that PRL does not regulate preadipocyte proliferation, but induces the expression of some early adipogenic markers. However, we believe that PRL alone is not sufficient for terminal differentiation, particularly for the accumulation of lipids. Finally, in strong agreement with this notion, it was recently reported that preadipocytes from PRLR -/- mice showed normal rates of proliferation and differentiation in vitro (unpublished observations by Flint, Binart, and Kelly) (6).

175 References

1. Hausman DB, DiGirolamo M, Bartness TJ, Hausman GJ, Martin RJ 2001 The biology of white adipocyte proliferation. Obes Rev 2:239-254 2. Prins JB, O'Rahilly S 1997 Regulation of adipose cell number in man. Clin Sci (Lond) 92:3-11 3. Faust IM, Johnson PR, Stern JS, Hirsch J 1978 Diet-induced adipocyte number increase in adult rats: a new model of obesity. Am J Physiol 235:E279-E286 4. Faust IM, Miller WH, Jr., Sclafani A, Aravich PF, Triscari J, Sullivan AC 1984 Diet- dependent hyperplastic growth of adipose tissue in hypothalamic obese rats. Am J Physiol 247:R1038-R1046 5. Oberbauer AM, Stern JS, Johnson PR, Horwitz BA, German JB, Phinney SD, Beermann DH, Pomp D, Murray JD 1997 Body composition of inactivated growth hormone (oMt1a- oGH) transgenic mice: generation of an obese phenotype. Growth Dev Aging 61:169-179 6. Flint DJ, Binart N, Kopchick J, Kelly P 2003 Effects of growth hormone and prolactin on adipose tissue development and function. Pituitary 6:97-102 7. Nanbu-Wakao R, Morikawa Y, Matsumura I, Masuho Y, Muramatsu MA, Senba E, Wakao H 2002 Stimulation of 3T3-L1 adipogenesis by signal transducer and activator of transcription 5. Molecular Endocrinology 16:1565-1576 8. Stephens JM, Morrison RF, Pilch PF 1996 The expression and regulation of STATs during 3T3-L1 adipocyte differentiation. Journal of Biological Chemistry 271:10441-10444 9. Ling C, Hellgren G, Gebre-Medhin M, Dillner K, Wennbo H, Carlsson B, Billig H 2000 Prolactin (PRL) receptor gene expression in mouse adipose tissue: increases during lactation and in PRL-transgenic mice. Endocrinology 141:3564-3572 10. McAveney KM, Gimble JM, Yu-Lee L 1996 Prolactin receptor expression during adipocyte differentiation of bone marrow stroma. Endocrinology 137:5723-5726 11. Viengchareun S, Bouzinba-Segard H, Laigneau JP, Zennaro MC, Kelly PA, Bado A, Lombes M, Binart N 2004 Prolactin potentiates insulin-stimulated leptin expression and release from differentiated brown adipocytes. J Mol Endocrinol 33:679-691 12. Pfaffl MW, Horgan GW, Dempfle L 2002 Relative expression software tool (REST) for group-wise comparison and statistical analysis of relative expression results in real-time PCR. Nucleic Acids Res 30:e36 13. Das R, Vonderhaar BK 1997 Prolactin as a mitogen in mammary cells. J Mammary Gland Biol Neoplasia 2:29-39 14. Van Coppenolle F, Skryma R, Ouadid-Ahidouch H, Slomianny C, Roudbaraki M, Delcourt P, Dewailly E, Humez S, Crepin A, Gourdou I, Djiane J, Bonnal JL, Mauroy B, Prevarskaya N 2004 Prolactin stimulates cell proliferation through a long form of prolactin receptor and K+ channel activation. Biochem J 377:569-578 15. Ortiz MB, Goin M, Gomez de Alzaga MB, Hammarstrom S, Jimenez dA 1995 Mevalonate dependency of the early cell cycle mitogenic response to epidermal growth factor and prostaglandin F2 alpha in Swiss mouse 3T3 cells. J Cell Physiol 162:139-146 16. Watanabe T, Satoh H, Togoh M, Taniguchi S, Hashimoto Y, Kurokawa K 1996 Positive and negative regulation of cell proliferation through prostaglandin receptors in NIH-3T3 cells. J Cell Physiol 169:401-409 17. de Asua LJ, Clingan D, Rudland PS 1975 Initiation of cell proliferation in cultured mouse fibroblasts by prostaglandin F2alpha. Proc Natl Acad Sci U S A 72:2724-2728

176 18. Nevalainen MT, Valve EM, Makela SI, Blauer M, Tuohimaa PJ, Harkonen PL 1991 Estrogen and prolactin regulation of rat dorsal and lateral prostate in organ culture. Endocrinology 129:612-622 19. Adachi H, Kurachi H, Homma H, Adachi K, Imai T, Morishige K, Matsuzawa Y, Miyake A 1994 Epidermal growth factor promotes adipogenesis of 3T3-L1 cell in vitro. Endocrinology 135:1824-1830 20. Bachmeier M, Loffler G 1995 Influence of growth factors on growth and differentiation of 3T3-L1 preadipocytes in serum-free conditions. Eur J Cell Biol 68:323-329 21. Smith PJ, Wise LS, Berkowitz R, Wan C, Rubin CS 1988 Insulin-like growth factor-I is an essential regulator of the differentiation of 3T3-L1 adipocytes. Journal of Biological Chemistry 263:9402-9408 22. Stewart WC, Baugh JE, Jr., Floyd ZE, Stephens JM 2004 STAT 5 activators can replace the requirement of FBS in the adipogenesis of 3T3-L1 cells. Biochem Biophys Res Commun 324:355-359 23. Nanbu-Wakao R, Fujitani Y, Masuho Y, Muramatu M, Wakao H 2000 Prolactin enhances CCAAT enhancer-binding protein-beta (C/EBP beta) and peroxisome proliferator- activated receptor gamma (PPAR gamma) messenger RNA expression and stimulates adipogenic conversion of NIH-3T3 cells. Molecular Endocrinology 14:307-316 24. Liu LH, Wang XK, Hu YD, Kang JL, Wang LL, Li S 2004 Effects of a fatty acid synthase inhibitor on adipocyte differentiation of mouse 3T3-L1 cells. Acta Pharmacol Sin 25:1052-1057 25. Hogan JC, Stephens JM 2005 The regulation of fatty acid synthase by STAT5A. Diabetes 54:1968-1975 26. Ling C, Svensson L, Oden B, Weijdegard B, Eden B, Eden S, Billig H 2003 Identification of functional prolactin (PRL) receptor gene expression: PRL inhibits lipoprotein lipase activity in human white adipose tissue. J Clin Endocrinol Metab 88:1804-1808

177 A 0.60

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Fig 1. 3T3-L1 preadipocyte viability curves. Cells were plated in 5% CSS at various densities and maintained for 24 hours before viability was measured by (A) MTT, (B), Resazurin, and (C) calcein AM assays. Data are expressed as optical density in A, and relative fluorescent units in B and C. Each value is a mean±SEM of 6 replicates from a representative experiment. Each experiment was repeated 2 times with similar results.

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Fig 2. Effects of 5% FBS or 5% CSS on 3T3-L1 growth curves. Cells (2000/well) were incubated in 5% FBS (red) or 5% CSS (blue) for up to 96 hours. Cell viability was assessed every 24 hours by (A) MTT, (B) resazurin, and (C) calcein AM assays. Data are expressed as optical density in A, and relative fluorescent units in B and C. Each value is a mean±SEM of 6 replicates from a representative experiment. Each experiment was repeated 2 times with similar results.

179 1.50 1.50

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Fig 3. Effect of PRL on 3T3-L1 preadipocyte growth, as determined by MTT. Cells (2000/well) were incubated in serum free media (SFM) (left column) or 1% CSS (right column) with increasing doses of PRL (0, 1, 5, 25, and 125 ng/ml) for 48 hours. 5% Fetal bovine serum (FBS) was used as a positive control. Data are expressed as optical density. Each value is a mean±SEM of 6 replicates from a representative experiment. Each experiment was repeated 3 times with similar results.

180 A 0.80 SFM 1.50 1% CSS

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Fig 4. Effect of PRL on 3T3-L1 preadipocyte growth, as determined by A) MTT, B) Rezasurin, and C) Calcein AM assays. Cells (2000/well) were incubated in serum free media (SFM) (left column) or 1% CSS (right column) with increasing doses of PRL (0, 1, 5, 25, and 125 ng/ml) for 48 hours. Fetal bovine serum (FBS) was used as a positive control. Data are expressed as optical density in A, and relative fluorescent units in B and C. Each value is a mean±SEM of 6 replicates from a representative experiment. Each experiment was repeated 4 times with similar results.

181 d0 d2 d4 d6 d8 Pit PRL

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4 PRLR expression 2 (Fold induction vs Day 0) 0 02468 Day of differentiation

Fig 5. PRL and PRLR expression during 3T3-L1 differentiation, as determined by conventional and real time PCR. Cells were grown to confluence and induced to differentiate in the presence of fetal bovine serum (upper panel) or charcoal/dextran treated (lower panel). Mouse pituitary (Pit) was used as a positive control, and was not expressed in 3T3-L1 cells. Real time values (lower panel) are the mean of three replicates and expressed as fold induction vs day 0.

182 1.00 FBS 0.80 CSS CSS+PRL 0.60

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Fig 6. Effect of PRL on lipid accumulation during 3T3-L1 adipogenesis. Cells were differentiated in the presence of 5% FBS (blue), 5% CSS (red), or 5% CSS+PRL (25 ng/ml) (purple) for 8 days. Lipids were stained with Oil Red O and extracted. Data are expressed as optical density. Each value is mean±SEM of 8 replicates. The experiment was performed 3 times with similar results.

183 Day 0

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Fig 7. Effect of PRL on lipid accumulation and morphological changes during adipogenesis in 3T3-L1 cells. Preadipocytes were differentiated in the presence of 5% FBS (left column), 5% CSS (middle column), or 5% CSS + PRL (25 ng/ml) (right column). Cells were fixed with paraformaldehyde throughout differentiation and stained with Oil Red O and hematoxylin, and then photographed.

184

Chapter 6. The Effects of PRL on Glucose Uptake in Mature Adipocytes

185 Introduction

Facilitated diffusion of glucose across the plasma membrane is mediated by a set of specific proteins called GLUTs (glucose trasporters) (1). These integral membrane proteins belong to an extended family of hexose transporters that determine the net rate of glucose entry into the cell. One member of this family, GLUT4, is expressed in adipose tissue where its activity is regulated by insulin. Binding of insulin to its receptor initiates a cascade of events involving phosphorylation of IRS proteins and the subsequent activation of PI3K. In turn, PI3 kinase activates downstream targets, such as Akt and PDK, both of which are involved in the translocation of GLUT4-containing vesicles to the plasma membrane, resulting in glucose uptake into the cell (2). Insulin also stimulates glucose uptake via two PI3K-independent pathways, the MAPK pathway and the CAP/Cbl pathway (3).

Under certain physiological conditions, PRL plays an important role in glucose metabolism. For example, during pregnancy, lactogenic hormones increase β-cell mass and insulin production (4). However, the effects of PRL on glucose transport are not known, with no reports of studies using adipocytes. In INS-1 cells, derived from rat islets, PRL increases GLUT2 expression, but has no effect in glucose-responsive MIN6 cells (5;6). In the lactating mammary gland, expression of GLUT4 decreases, while GLUT1 increases, suggesting that PRL may play a role in the regulation of glucose uptake by altering transporter levels (7).

PRL activates its cognate receptor, which dimerizes and activates Jak2. The activated Jak2 mediates phosphorylation of various signaling molecules, primarily the Stats, but also IRS-1, and IRS-2 (8). Therefore, PRLR-mediated phosphorylation of IRS-1 by Jak2 could exert some insulin-like effects, including glucose transport. Other cytokines, including leptin induce glucose uptake in a human neuronal cell line via a PI3K-dependent mechanism (9). However, few studies have investigated the effects of cytokines on glucose uptake in adipocytes. Therefore, the purpose of this set of studies was to determine the effects of PRL on basal and insulin-stimulated glucose uptake in differentiated 3T3-L1 adipocytes.

Materials and methods

Reagents

3T3-L1 cells were obtained from Dr. Philip Scherer, Albert Einstein College of Medicine. Trypsin, and high- and low-glucose Dulbecco’s modified Eagle medium (DMEM) were purchased from Gibco (Carlsbad, CA). Fetal bovine serum and Charcoal stripped serum (CSS) were purchased from JRH biosciences (Lenexa, KA) and Hyclone (Logan, UT), respectively. Dexamethasone, 1-methyl-3-isobutylxanthine (IBMX), bovine insulin, and sodium dodecyl sulfate (SDS) were obtained from Sigma-Aldrich (St. Louis, MO). [3H]-2-deoxyglucose was purchased from American Radiolabeled Chemicals (St. Louis, MO). Microscint cocktail was purchased from Perkin-Elmer. Ovine PRL was a gift from Dr. Arieh Gertler (The Hebrew University of Jerusalem).

Cell culture

186 3T3-L1 preadipocytes were maintained at 37°C in low glucose DMEM and 5% FBS in a 5% CO2 environment. Cells were split every two days to prevent them from reaching confluence. For experiments, cells were plated at 11,000 cells/cm2 and grown to confluence in low-glucose DMEM and 5% FBS. Two days after reaching confluence, cells were induced to differentiate by changing the medium to high-glucose DMEM containing 5% fetal bovine serum, 0.5 mM 3- isobutyl-methylxanthine, 0.5 µM dexamethasone, and 10μg/ml insulin. After 48 h, this medium was replaced with DMEM supplemented with 5% fetal bovine serum, which was changed every 2 days until the cells were used for experimentation. Experiments were only performed on fully differentiated 3T3-L1 adipocytes.

Glucose uptake

3T3-L1 preadipocytes were differentiated in 48 well plates in the presence of FBS for 10-12 days as described in chapter 5. For experiments, cells were rinsed 3 times with SFM, then starved in either serum-free media (SFM) or 3% CSS for 24 hours. Adipocytes were rinsed another 3 times with SFM and then incubated in low-glucose DMEM containing 1% CSS and oPRL (0, 1, 5, 25, 125 ng/ml) for 10 minutes to 24 hours. After rinsing again, cells were incubated at 37°C for 20 minutes in 270 μl of Fat Cell Buffer (FCB) (Kreb’s Ringer w/ 25 mM Hepes, 125 mM NaCl, 5 4 mM KCl, 1.8 mM CaCl2, 2.6 mM MgSO w/ 2 mM Pyruvate) either with or without insulin (100nm). Next, 30μl of PBS containing 0.5 μCi of [3H] 2-deoxyglucose and 1.1 mM cold-2- deoxyglucose was added for 20 minutes. Cells were placed on ice and washed 4 times with cold PBS. Following cell lysis 2% SDS, 100μl of cell lysate was removed for determination of incorporated radioactivity by scintillation counting using a Top Count (Beckman-Coulter), and two 5 μl aliquots were used for protein determination by BCA analysis (10). Data are expressed as CPM/μg of protein.

GLUT4 expression by real time PCR

Cells were differentiated in 6 well plates for 10-12 days as above. After overnight serum starvation, cells were rinsed 3 times with SFM, and then incubated with oPRL (0, 1, 5, 25, 125 ng/ml) for 8 or 24 hours. Total RNA was extracted from using Tri Reagent (MRC, Cincinnati, OH), and 5 μg of RNA was reverse transcribed as described in chapter 4. PCR was performed on 200 ng of cDNA using intron-spanning primers for GLUT4 and β-actin, as described in Chapter 4. β-Actin was used as a reference gene. Fold changes in gene expression were calculated from the cycle threshold measurements, using the method of Pfaffl et al. (11).

Data Analysis

For real time PCR, gene expression was analyzed using REST (relative expression software tool V1.9.9) (11). Statistical differences were determined by one-way ANOVA followed by post hoc analysis comparing least significant differences. P values <0.05 were considered significant.

Results

PRL does not have consistent effects on glucose uptake in mature 3T3-L1 adipocytes

187 To determine the effects of PRL on glucose uptake in adipocytes, differentiated 3T3-L1 cells were pretreated with PRL, followed by measurement of basal or insulin-stimulated glucose incorporation. As shown in Fig. 1, incubation with PRL for 24 hr had inconsistent effects on glucose uptake. In the left panel, which is representative of 3 experiments, 5 ng/ml and 25 ng/ml of PRL induced a 4-5 fold increase in basal glucose incorporation compared to non-treated cells. Insulin alone increased glucose uptake by ~4-fold, and both 5 ng/ml and 25 ng/ml PRL pretreatment doubled this effect. However, in the right panel, also representative of 3 experiments, PRL treatment had no effect on basal glucose uptake. Insulin increased glucose incorporation by ~4-fold, while PRL pretreatment had no effect.

As shown in Fig. 2, short-term PRL treatment also had no effect on glucose uptake in these cells. Cells were either treat with PRL for 1 hour (left panel), or 10 minutes (right panel) prior to the addition of radio-labeled glucose. Neither treatment had any effect on basal or insulin-stimulated glucose uptake. In all experiments, insulin alone increased glucose transport by 4-6-fold.

GLUT4 expression

To determine the effects of PRL on GLUT4 expression, differentiated cells were treated for 24 hours, and Real Time PCR was used to measure mRNA levels. As shown in Fig 3, PRL induced variable changes in GLUT4 expression. In the left panel, which is representative of 2 experiments, GLUT4 mRNA increased approximately 1-fold when treated with 1 ng/ml PRL, while higher doses (5 and 25 ng/ml) increased expression by ~2-3 fold. As evident in the right panel, the same treatment had no effect on GLUT4 levels. Similar results were seen in 3 separate experiments.

Discussion

This report describes inconsistent effects of PRL on glucose transporter activity and expression in 3T3-L1 adipocytes. In some experiments, long-term PRL treatment did elicit a response on GLUT4 expression, while increasing basal and insulin-stimulated rates of glucose incorporation. Short-term PRL treatment had no effect on either basal or stimulated glucose transport. Unfortunately, the inability to consistently reproduce these results has prevented us from thoroughly investigating this phenomenon.

The observation that chronic PRL treatment alters glucose transport is not surprising. There are several reports that members of the cytokine family are capable of long-term regulation of glucose uptake. For example, chronic treatment of 3T3-L1 adipocytes with superphysiological concentrations of GH (0.5 μg/ml) decreased glucose uptake, despite enhanced tyrosine phosphorylation of IRS-1 and its association and activation of PI3K (12). This effect is proposed to be a result of uncoupling between activation of PI3K and its downstream signals. The downstream targets involved in insulin signaling have yet to be clearly identified, and the mechanism by which GH exerts this effect is unknown. In our experiments, chronic PRL exposure had the opposite effect, increasing glucose transport. Therefore, we attempted to elucidate the underlying mechanism.

188 There are few reports of PRL-mediated alterations of glucose uptake. In the mammary gland, PRL increases lactose synthesis by inducing glucose transport (13). Although the underlying mechanism was not investigated, it is possible that PRL modulates insulin-binding capacities in the mammary gland, as reported by others (14). While this could be a possible explanation, we observed an increase in both basal and insulin-stimulated glucose transport. Moreover, The PRL- mediated induction of glucose transport appears to be additive, with nearly the same increases observed under both basal and insulin-stimulated conditions. To determine the mechanism by which PRL causes this increase, we measured mRNA levels of GLUT4, the primary regulator of glucose absorption in adipocytes. In several of our experiments, chronic PRL exposure indeed elevated GLUT4 levels. GLUT4 is an insulin responsive transporter, which is constantly cycling to the plasma membrane, contributing the basal rate of glucose uptake. Therefore, increased presence of GLUT4 in the plasma membrane under non-stimulated conditions could certainly result in an elevated rate of basal glucose uptake. Furthermore, a higher number of glucose transporters in cycling endosomes may similarly augment the insulin-stimulated glucose response. Indeed, chronic exposure to TNF-α decreases glucose uptake in 3T3-L1 adipocytes by down-regulating the expression of GLUT4 (15). Moreover, long-term exposure of neurons to leptin down-regulates GLUT4 levels and abolishes the acute stimulation of glucose uptake in response to insulin treatment (9). Increased expression of GLUT4 may not be sufficient to increase glucose uptake. Indeed, in 3T3-L1 adipocytes, the gp160 cytokine ciliary neurotrophic factor (CNTF), has no effect on glucose uptake despite increasing expression of GLUT4 (16).

There are several reports of acute cytokine-mediated changes in glucose transport. In one study a five minute treatment with high concentrations of GH (0.5 μg/ml) increased glucose uptake and GLUT4 translocation to the plasma membrane in 3T3-L1 cells via a PI3K-independent mechanism (17). While we did not observe short-term induction of glucose transport by PRL, the doses used in our study were relatively low, but well within the non-lactating physiological range. Therefore, the amount of PRL used in our experiments may have been sufficient enough to activate signaling pathways that are not involved in glucose uptake, but not enough to activate those that are necessary for glucose uptake. Indeed, in 3T3-L1 cells, PRL, as well as lower concentrations of GH (125 ng/ml) primarily activate the Stat5 pathway. However, a recent study demonstrated that Stat 5 proteins are not mediators of insulin action in these cells (18).

In conclusion, our studies indicate that chronic PRL treatment may increase glucose uptake in adipocytes, possibly via up-regulation of GLUT4 expression, whereas acute PRL treatment exerted no effects. We repeated these studies under a number of different experimental conditions, but unfortunately were unable to consistently reproduce our positive data. In addition, several experiments were performed using cultured adipose tissue explants and isolated adipocytes. However, due to the repeated washes and manipulations in this experimental procedure, primary adipocytes unavoidably became damaged and unusable.

189

References

1. Joost HG, Thorens B 2001 The extended GLUT-family of sugar/polyol transport facilitators: nomenclature, sequence characteristics, and potential function of its novel members (review). Mol Membr Biol 18:247-256 2. Watson RT, Pessin JE 2006 Bridging the GAP between insulin signaling and GLUT4 translocation. Trends Biochem Sci 31:215-222 3. Baumann CA, Ribon V, Kanzaki M, Thurmond DC, Mora S, Shigematsu S, Bickel PE, Pessin JE, Saltiel AR 2000 CAP defines a second signalling pathway required for insulin- stimulated glucose transport. Nature 407:202-207 4. Sorenson RL, Brelje TC, Roth C 1993 Effects of steroid and lactogenic hormones on islets of Langerhans: a new hypothesis for the role of pregnancy steroids in the adaptation of islets to pregnancy. Endocrinology 133:2227-2234 5. Petryk A, Fleenor D, Driscoll P, Freemark M 2000 Prolactin induction of insulin gene expression: the roles of glucose and glucose transporter-2. Journal of Endocrinology 164:277-286 6. Shao J, Qiao L, Friedman JE 2004 Prolactin, progesterone, and dexamethasone coordinately and adversely regulate glucokinase and cAMP/PDE cascades in MIN6 beta- cells. Am J Physiol Endocrinol Metab 286:E304-E310 7. Burnol AF, Leturque A, Loizeau M, Postic C, Girard J 1990 Glucose transporter expression in rat mammary gland. Biochem J 270:277-279 8. Goffin V, Binart N, Touraine P, Kelly PA 2002 Prolactin: the new biology of an old hormone. Annu Rev Physiol 64:47-67 9. Benomar Y, Naour N, Aubourg A, Bailleux V, Gertler A, Djiane J, Guerre-Millo M, Taouis M 2006 Insulin and leptin induce Glut4 plasma membrane translocation and glucose uptake in a human neuronal cell line by a phosphatidylinositol 3-kinase- dependent mechanism. Endocrinology 147:2550-2556 10. LOWRY OH, ROSEBROUGH NJ, FARR AL, RANDALL RJ 1951 Protein measurement with the Folin phenol reagent. Journal of Biological Chemistry 193:265- 275 11. Pfaffl MW, Horgan GW, Dempfle L 2002 Relative expression software tool (REST) for group-wise comparison and statistical analysis of relative expression results in real-time PCR. Nucleic Acids Res 30:e36 12. Takano A, Haruta T, Iwata M, Usui I, Uno T, Kawahara J, Ueno E, Sasaoka T, Kobayashi M 2001 Growth hormone induces cellular insulin resistance by uncoupling phosphatidylinositol 3-kinase and its downstream signals in 3T3-L1 adipocytes. Diabetes 50:1891-1900 13. Oppat CA, Rillema JA 1988 Characteristics of the early effect of prolactin on lactose biosynthesis in mouse mammary gland explants. Proceedings of the Society for experimental Biology and Medicine 188:342-345 14. Vernon RG, Flint DJ 1983 Control of fatty acid synthesis in lactation. Proc Nutr Soc 42:315-331 15. Stephens JM, Pekala PH 1991 Transcriptional repression of the GLUT4 and C/EBP genes in 3T3-L1 adipocytes by tumor necrosis factor-alpha. Journal of Biological Chemistry 266:21839-21845

190 16. Zvonic S, Cornelius P, Stewart WC, Mynatt RL, Stephens JM 2003 The regulation and activation of ciliary neurotrophic factor signaling proteins in adipocytes. Journal of Biological Chemistry 278:2228-2235 17. Sakaue H, Ogawa W, Takata M, Kuroda S, Kotani K, Matsumoto M, Sakaue M, Nishio S, Ueno H, Kasuga M 1997 Phosphoinositide 3-kinase is required for insulin-induced but not for growth hormone- or hyperosmolarity-induced glucose uptake in 3T3-L1 adipocytes. Molecular Endocrinology 11:1552-1562 18. Zvonic S, Story DJ, Stephens JM, Mynatt RL 2003 Growth hormone, but not insulin, activates STAT5 proteins in adipocytes in vitro and in vivo. Biochem Biophys Res Commun 302:359-362

191

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Fig 1. Chronic PRL treatment does not have a consistent effect on glucose uptake in 3T3-L1 adipocytes. Cells were treated with PRL for 24 hours, then the rate of basal and 10 minute insulin-stimulated 3H-2-deoxyglucose were measured. Left and right panels represent the results obtained from two different experiments Data are expressed as CPM/μgof protein. Each value is a mean±SEM of 4 replicates. The experiment was repeated 6 times with similar variable results.

192 500 400

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Fig 2. Short term PRL treatment has no effect on glucose uptake in 3T3-L1 adipocytes. Cells were treated with PRL for 10 minutes (left panel) or 1 hour (right panel), then the rate of basal and 10 minute insulin-stimulated 3H-2-deoxyglucose were measured. Data are expressed as CPM/μg of protein. Each value is a mean±SEM of 4 replicates. The experiment was repeated 3 times with similar results.

193 3 3

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Fig 3. 24 hour PRL treatment does not have a consistent effect on GLUT4 expresion in 3T3- L1 adipocytes, as determined by real time PCR. Left and right panels represent the results obtained from two different experiments. Data are expressed as fold-induction vs control treatment. Each value is a mean±SEM of 3 replicates. The experiment was repeated 4 times with similar variable results.

194

Chapter 7. General Conclusions

195 The functions of PRL during the estrous cycle, pregnancy and lactation have been investigated extensively. However, far less attention has been paid to its effects in males and non-lactating females. Given the recent reports implicating PRL in metabolic homeostasis, we hypothesized that PRL is required for normal weight gain and adiposity in male and non-lactating female mice. We postulated that this may occur by PRL-mediated alterations in adipokine secretion, glucose metabolism, and/or adipose tissue growth and metabolism. The studies conducted in this thesis showed that at non-lactating levels, the contribution of PRL to metabolic homeostasis in mice is negligible. It would be easy to conclude that under these physiological conditions PRL is not an important metabolic hormone. However, extrapolating data from mice to other species should be done with caution.

Several important considerations should be made when examining interspecies differences in the functions of PRL. The PRL molecule is defined by its general structure, especially the location of disulphide bridges. While the sequence homology between humans and rodents is ~60%, there are some 32 residues conserved among different species (1). These cluster in distinct regions of the molecule considered to be determinants for receptor binding and activation. Given the structural and organizational similarities between PRL molecules, we must question how PRL regulates such a myriad of functions across species, as well as within one organism. The pleiotropic actions of PRL are undoubtedly due to the presence of PRLRs on multiple cell types, as well as the ability of PRL to activate interacting signaling pathways. PRL also undergoes post- translational modifications, including glycosylation, phosphorylation, and proteolytic cleavage, which generates several variants. Notably, the types and extent of modifications differ among species and likely contribute to the pleiotropic functions of PRL.

The sites of PRL production vary considerably between species. In humans, PRL is produced not only by the pituitary, but by many extrapituitary tissues, while this is not the case in rodents (2). Although the concentrations of locally produced PRL are difficult to evaluate, extrapituitary PRL must be considered as an autocrine/paracrine regulator of metabolism. Notably, this may be particularly relevant to human adipose tissue, which produces PRL and expresses its receptor. Since the relative contributions of circulating and adipose derived PRL in humans are unknown, hyperprolactinemia in humans does not necessarily reflect the situation at the level of the adipocyte.

Notably, there are no known cases of a non-functional or an absent PRLR in humans. A mutation in PRL or the PRLR that disrupts receptor activation or signaling could result in no obvious phenotype. This would indicate either that PRL is not important in human metabolism, or that compensatory mechanisms exist. On the other hand, PRL may be so important in humans that a mutation that disrupts PRLR activation causes death in utero. At this point, it is uncertain which is the case in humans, but it is important to recognize that the human fetus, uniquely among species, is exposed to very high concentrations of PRL that is produced by the decidua.

Although studies with transgenic mice have yielded a wealth of knowledge, the substantial differences in metabolic homeostasis between rodents and humans must be taken into account. For example, unlike rodents, circulating leptin levels do not change acutely after eating in humans, and the promise of leptin as an anti-obesity treatment in rodents has not materialized to human therapy. An interspecies difference is also apparent in in vitro systems. GH decreases

196 proliferation of murine 3T3-F442A preadipocytes but stimulates proliferation in primary human preadipocytes (3). Furthermore, serum is required for efficient adipogenesis in 3T3-L1 cells, while human preadipocytes undergo differentiation without serum, with BSA actually inhibiting their differentiation (4).

Indeed, PRL regulates hundreds of functions among species. This is well exemplified by the osmoregulatory role of PRL in fish, its pronounced effects on hair follicles in sheep, and its unique control of corpus luteum function in rodents. It is not totally surprising, therefore, that PRL appears to have metabolic functions in some species, but not in mice.

Summary of my studies

My studies on the metabolic functions of PRL began by measuring the expression of selected genes in a small number of wild-type and PRL-knockout mice. The initial results were exciting and strongly suggested a role for PRL in metabolism. We were cautious in our approach though, since these experiments compared gene expression in tissues isolated from commercially purchased wild-type mice to those in PRL-deficient mice maintained in Dr. Horseman’s laboratory. To properly study this phenomenon, it was necessary to increase the number of mice, compare littermates, and include heterozygous mice as an additional experimental group. Indeed, data can be misconstrued based on an insufficient number of observations, which we suspect was the case in at least one report of reduced body weight in PRLR-deficient mice. Therefore, we were careful to include a large number of animals/replicates to ensure that our data were accurate.

In order to conduct our studies, we established a mouse colony in our facility, a process which required significant time for animal quarantine, transfer, and breeding. Although the initial differences in gene expression were not sustained when littermates were compared, some of our other preliminary data were promising, such as differential body composition profiles in male wild-type and PRL-deficient mice. To properly examine this observation, we expanded our study to include more animals, and transferred a portion of our colony to another facility where weight gain and adiposity were measured for several months. Thus, a great deal of time was spent on colony maintenance and data collection in two locations.

Our in vivo studies extended beyond measurement of gross metabolic parameters, and indicate that PRL regulates other aspects of metabolic homeostasis, particularly adipokine release. Indeed, leptin levels were altered by PRL-deficiency under certain physiological conditions. Since fat content was unchanged in these animals, we do not believe that leptin levels were merely a reflection of adiposity. Because of a strong association between leptin and energy balance, we initially planned to compare food intake and energy expenditure between wild-type and knockout mice. However, since the differences in leptin were seen under only one physiological condition, we decided that performing these experiments would be an impractical use of valuable time and resources.

Although PRLR-deficient mice have been used to study the metabolic actions of PRL, a ligand- specific knockout mouse model, such as the PRL-deficient mouse, offers many advantages. Notably, the presence of an intact PRLR signaling mechanism enables application of PRL

197 replacement therapy. In fact, while conducting our in vivo studies, we developed a technique for PRL replacement using permeable hollow fibers filled with lactotrophs. The enclosed cells survived well in vitro and produced significant amounts of PRL for at least 12 weeks. However, given that our studies yielded mostly negative data, we decided that it was not an avenue worth pursuing.

Because adipose tissue from mice provides an insufficient number of preadipocytes to perform our studies, we used murine 3T3-L1 preadipocytes as a model. Certainly, the optimal method for studying these phenomena in the absence of PRL signaling would be via blockade of the PRLR. However, given the lack of effective PRLR antagonists, we employed an additive strategy, supplementing CSS with exogenous PRL. Because we wished to determine the metabolic functions of PRL under non-lactating conditions, we used PRL concentrations less than 250 ng/ml. Some of our in vitro experiments also yielded initial promising results. We repeated these experiments several times, and given the length of time required to differentiate preadipocytes, we spent a considerable amount of time and effort trying to reproduce these results under many conditions and treatment paradigms.

Given the uncertainty of the metabolic functions of PRL, we maintain that these studies were necessary. In fact, the in vivo data were peer reviewed and selected for publication in a highly regarded scientific journal, acknowledging the controversy over whether PRL is an important metabolic hormone in mice. We carefully examined numerous metabolic parameters with which PRL has been associated, and expanded our studies to investigate new factors such as FIAF. The use of gene array or proteomics should prove useful for analyzing a large number of metabolic parameters simultaneously.

Our in vitro experiments yielded results that are not likely to be submitted for publication. However, we feel that our experimental design was appropriate and our data are valid. Further supporting the notion that PRL has minor metabolic actions in mice, additional studies performed in our lab using murine 3T3-F442A adipocytes have found no clear function for PRL. On the other hand, studies with our lab’s newly generated LS14 human adipocytes show that PRL inhibits lipolysis as well as the secretion leptin, adiponectin, and IL-6. Hence, further studies on the metabolic actions of PRL should be performed with human adipocytes. In conclusion, more than most hormones, the physiological actions of PRL are species specific, and perhaps mice are not a good model for studying the metabolic actions of PRL.

Finally, I greatly benefited from my graduate school experience in several ways. First, as part of investigating such a controversial topic, it was necessary that I comprehend, interpret, and develop my own opinion based on a large body of often contradictory literature. Second, I was permitted to decide which hypotheses warranted investigation. Since these studies explored a wide variety of metabolic parameters, I learned and performed a considerable number of experimental techniques, both in vivo and in vitro. In conclusion, this experience served as an excellent foundation to continue studying factors that regulate metabolism, perhaps with an emphasis on adipocytes.

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References

1. Sinha YN 1995 Structural variants of prolactin: occurrence and physiological significance. Endocr Rev 16:354-369 2. Ben Jonathan N, Mershon JL, Allen DL, Steinmetz RW 1996 Extrapituitary prolactin: distribution, regulation, functions, and clinical aspects. Endocr Rev 17:639-669 3. Flint DJ, Binart N, Kopchick J, Kelly P 2003 Effects of growth hormone and prolactin on adipose tissue development and function. Pituitary 6:97-102 4. Schlesinger JB, van H, V, Alberti-Huber CE, Hauner H 2006 Albumin inhibits adipogenesis and stimulates cytokine release from human adipocytes. Am J Physiol Cell Physiol 291:C27-C33

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Chapter 8. Curriculum Vitae

200 Christopher Ryan LaPensee

Address: Home: 3221 Grischy Lane Cincinnati OH 45208

Work: Neuroscience Graduate Program University of Cincinnati 3125 Eden Avenue Cincinnati, OH 45267-0521

Phone: 513-310-1337 (home) 513-558-4824 (laboratory) Fax: 513-558-4823 Email: [email protected]

Education

2000 B.S. University of Michigan, Ann Arbor, MI. May 2000 Major: Biochemistry Advisor: Dr. David L Turner, Department of Neurology

2000-present: Graduate Student, Neuroscience Graduate Program, University of Cincinnati Thesis Advisor: Dr. Nira Ben-Jonathan, Department of Cell Biology

Professional Experience

1997-1998 Undergraduate Research Assistant, Dr Robert Macdonald Department of Neurology, University of Michigan, Ann Arbor

Research Assistant at the University of Michigan School of Nursing, Collect data on dementia patients in nursing homes

EKG Technician at the University of Michigan Hospital

Blood/Gas Lab Technician at the University of Michigan Hospital Hematology Lab

1999-2000 Undergraduate Research Assistant, Dr. David L Turner Department of Neurology, University of Michigan, Ann Arbor

2001 Research Rotation, Dr. Nancy Ratner Department of Cell Biology, University of Cincinnati

2000-Present: Graduate Research Assistant, Dr. Nira Ben-Jonathan

201 Department of Cell Biology, University of Cincinnati

Awards

2000 NSF Summer undergraduate training grant in developmental neurobiology, University of Michigan Department of Neurology 2000 NIH Predoctoral neuroscience training grant, University of Cincinnati, 2 years training period

2002 NIH training grant in Developmental and Perinatal Endocrinology, University of Cincinnati, Children’s Hospital Medical Center

2004 Travel Award for the 86th Annual Meeting of the Endocrine Society, New Orleans, Louisiana

Research Interests

Hormonal regulation of metabolic homeostasis in mice: Investigating the effects of prolactin on body weight and fat composition, lipid profile, glucose metabolism, and the expression of adipose-specific genes in male prolactin-knockout mice.

Effects of prolactin on preadipocyte proliferation, differentiation and metabolism: Investigating the mitogenic and adipogenic effects of PRL in 3T3-L1 preadipocytes.

Application of proteomics and gene arrays for studying the effects of prolactin on adipokine expression and release: Examine the effect of PRL on global production and release of cytokine/adipokines using LS14 human adipocytes as the cellular model.

Publications

Ben-Jonathan N, Hugo E.R., Brandebourg T.D., LaPensee C.R. Focus on Prolactin as a Metabolic Hormone. Trends in Endocrinology and Metabolism 17:110-116, 2006.

LaPensee C.R., Horseman N.D., Tso P, Brandebourg T.D., Hugo E.R., Ben-Jonathan N. The prolactin-deficient mouse has an unaltered metabolic phenotype. Endocrinology 147: 4638- 4645, 2006.

Ben-Jonathan N, LaPensee C.R. Prolactin and its neuroendocrine control, in: Encyclopedia of Neuroscience, Squire, L.R. et al (eds), Elsevier, Oxford, UK (in press).

Bigsby R.M., LaPensee C.R., Ben-Jonathan N, Caperelle-Grant A. Prolactin mediates estrogen- induced cell proliferation in the uterine stroma and myometrium following progesterone priming (in preparation).

202 Fisher K., LaPensee C.R., Janik J., Reinscheid R.K., Murphree E., Ben-Jonathan N., Callahan P. Orphanin FQ/nociceptin gene deletion decreases prolactin secretion and pup survival in post-partum mice (in preparation).

Memberships

2002-present Associate member, The Endocrine Society

Laboratory Techniques

Transgenic animal husbandry and genotyping Small animal surgery Cell/tissue culture RealTime PCR Western blotting Immunohistochemistry Gene cloning Cell proliferation, apoptosis and viability assays Nb2 bioassay for prolactin Radioactive glucose uptake

Basic skills in programming, proficient in Excel, Sigma plot, and SlideWrite

Teaching/Outreach experience

1998-1999 Organic chemistry peer tutor, University of Michigan Volunteer: University of Michigan Hospitals Central Pharmacy Volunteer: University of Michigan Geriatrics Center, Silver Club, a program specifically designed to meet the needs of older adults with dementia/memory loss

Conference Attendance

2001 83rd Annual Meeting of the Endocrine Society, Denver, Colorado 2002 84th Annual Meeting of the Endocrine Society, San Francisco, California 2003 85th Annual Meeting of the Endocrine Society, Philadelphia, Pennsylvania 2004 Gordon Conference: Prolactin Family, Ventura, California 2004 86th Annual Meeting of the Endocrine Society, New Orleans, Louisiana 2005 87th Annual Meeting of the Endocrine Society, San Diego, California 2006 88th Annual Meeting of the Endocrine Society, Boston, Massachusetts

Presentations

2003 Neuroscience Symposia, Prolactin production by Adipose Tissue, University of Cincinnati

203 Neuroscience Seminar Series, The Role of Prolactin in Preadipocyte Proliferation and Differentiation, University of Cincinnati

2004 Neuroscience Seminar Series, Prolactin as a Regulator of Adipose Tissue Homeostasis in Mice, University of Cincinnati Poster Presentation, Differential Expression of Adipose-Specific Genes in Prolactin Knockout Mice, Gordon Conference: Prolactin Family, Ventura, CA Poster Presentation, Differential Expression of Adipose-Specific Genes in Prolactin Knockout Mice, 86th Annual Meeting of the Endocrine Society, New Orleans, LA

2005 Neuroscience Seminar Series, The Role of Prolactin in Adipose Tissue Homeostasis, University of Cincinnati Poster Presentation, The Effects of Prolactin on 3T3-L1 Proliferation and Metabolism, Graduate Student Research Forum, University of Cincinnati

2006 Neuroscience Seminar Series, The Prolactin-Deficient Mouse has an Unaltered Metabolic Phenotype Poster Presentation, The Prolactin-Deficient Mouse has an Unaltered Metabolic Phenotype, 88th Annual Meeting of the Endocrine Society, Boston, MA

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