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Treatment of obesity and diabetes by a regulatable leptin cell therapy system Oosman, Sarah Natasha 2005

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TREATMENT OF OBESITY AND DIABETES BY A REGULATABLE LEPTIN CELL THERAPY SYSTEM by SARAH NATASHA OOSMAN B.Sc. (Phyl.), The University of Saskatchewan, 1995 B.Sc. (PT), The University of Saskatchewan, 1998 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES (Physiology) THE UNIVERSITY OF BRITISH COLUMBIA May 2005 © Sarah N. Oosman, 2005 u ABSTRACT Obesity, a chronic disorder that is increasing in prevalence worldwide, is a major risk factor for the development of type 2 diabetes, a metabolic disorder characterized by hyperglycemia. In this thesis, the efficacy of a leptin cell therapy was evaluated for the treatment of obesity and diabetes in leptin-deficient ob/ob mice and in high fat fed, leptin-resistant C57BL/6J mice. A gut endocrine K-cell line was engineered to produce leptin under the regulation of an RU486 controlled GeneSwitch™ system. In culture, these cells express and release leptin in an RU486 dose- and time-dependent manner. These cells were transplanted into ob/ob mice via (a) kidney capsule and (b) intraperitoneal (IP) injection of alginate encapsulated cells along with 14-day RU486 pellets. In mice that received leptin-producing cells under the kidney capsule, reductions in body weight (10%), food intake (50%) and blood glucose levels (67%) were observed 10 days post-transpjant, relative to controls. Body weight and food intake rapidly increased thereafter to that of controls. Interestingly, however, blood glucose concentrations remained reduced by 40% up to 2 weeks post-transplantation. Animals given IP encapsulated cells lost up to 17% of body weight and then rapidly returned to their starting weight 14 days later at exhaustion of the RU486. Remarkably, despite the fact that body weight was completely regained within 20 days post-transplantation, blood glucose concentrations remained reduced by almost 70% up to 50 days post-transplantation. Both the number of transplanted cells and the dose of RU486 given could regulate the effects of leptin cell therapy. Obese C57BL /6J mice on a high fat diet did not respond with reductions in body weight, food intake or blood glucose levels after being transplanted with encapsulated leptin-producing gut cells. These data demonstrate that leptin administered via a cell therapy strategy can result in a reduction in body weight, food intake and long term corrections of blood glucose Ill concentrations in leptin sensitive ob/ob mice but not in high fat fed leptin resistant C57BL/6J mice under the conditions tested. iv TABLE OF CONTENTS Page ABSTRACT ii TABLE OF CONTENTS iv LIST OF FIGURES vii LIST OF TABLES x LIST OF ABBREVIATIONS xi ACKNOWLEDGEMENTS xii INTRODUCTION 1 Obesity and Type 2 Diabetes 1 Parabiosis experiments and the sateity signal 3 Leptin 4 Leptin receptor 7 Leptin in the treatment of obesity and diabetes 8 Leptin cell therapy 11 Transplantation strategies 14 THESIS INVESTIGATION 16 MATERIALS AND METHODS 17 The GeneSwitch™ System 17 Generation of the control gut cell line and the leptin-producing gut cell line using the GeneSwitch™ System 19 In vitro studies 25 Northern blot determination of GIP mRNA expression 25 Effect ofRU486 dose and exposure time on leptin mRNA expression in leptin-producing gut cells 26 V Western blot analysis of leptin protein expression in leptin-producing gut cells 26 ELISA analysis of leptin release from leptin-producing gut cells 27 Encapsulation of leptin-producing or control gut cells 28 RU486 induction of encapsulated versus non-encapsulated leptin-producing gut 29 cells RU486 induction of encapsulated leptin-producing gut cells cultured for 5 weeks 29 Animals 30 RU486 pellets 31 In vivo studies 34 Transplantation of leptin-producing gut cells under the kidney capsule 35 Transplantation of encapsulated leptin-producing gut cells or control gut cells 36 DATA A N A L Y S I S 41 R E S U L T S 42 S T U D Y I: In vitro characterization of leptin-producing gut cells 42 S T U D Y II: Treatment of obesity and diabetes in ob/ob mice by transplantation of leptin-producing gut cells under the kidney capsule 49 S T U D Y III: Treatment of obesity and diabetes in ob/ob mice by transplantation of encapsulated leptin-producing gut cells 61 S T U D Y IV: Treatment of obesity and diabetes in high-fat fed C57BL /6J mice by transplantation of encapsulated leptin-producing gut cells 101 D I S C U S S I O N 123 Regulation of leptin m R N A and protein expression and secretion from leptin-producing gut cells 123 Treatment of diabetes and obesity in ob/ob mice by leptin cell therapy 126 Leptin cell therapy in the treatment of obesity and diabetes in high-fat fed C57BL /6J 135 mice CONCLUSIONS REFERENCES V l l LIST OF FIGURES Figure 1 Schematic representation of the GeneSwitch™ System transfected into GTC-1 cells 18 Figure 2 A Schematic representation of the steps involved in construction the pSwitch GIPro plasmid 21 Figure 2 B Schematic representation of the steps involved in construction the pGemB/hLeptin/-V5His plasmid 23 Figure 3 GIP m R N A expression in GTC-1 cells compared to control gut cells (GTC-1 pSwitchGIPro cells) and leptin-producing gut cells (GTC-1 pSwitchGIProhLeptin cells) 43 Figure 4 Leptin m R N A expression in leptin-producing gut cells 46 Figure 5 Leptin protein expression in leptin-producing gut cells 47 Figure 6 Leptin protein release from leptin-producing gut cells 48 Figure 7 Transplantation of control gut or leptin-producing gut cells under the kidney capsule of ob/ob mice 51 Figure 8 Human leptin radioimmunoassay: comparison of results obtained from an A L P C O human leptin RIA (A) and a LINCO human leptin RIA (B) 54 Figure 9 Kidney capsule transplantation of control gut or leptin-producing gut cells into ob/ob mice, with or without Cyclosporine (Cs) 57 Figure 10 Serum human leptin levels from ob/ob mice that received leptin cell therapy via kidney capsule transplant, with or without Cyclosporine (Cs) 59 Figure 11 Photograph of an ob/ob mouse that received leptin-producing gut cells to the kidney capsule with Cycosporine (Cs) treatment 60 Figure 12 Encapsulation of cells in a sodium-alginate polymer coating 62 Figure 13 Leptin protein secretion from (A) non-encapsulated or (B) encapsulated leptin-producing gut cells 63 Figure 14 The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose in ob/ob mice 66 Figure 15 The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose in ob/ob mice transplanted with various volumes of cells 70 Figure 16 The effect of leptin cell therapy on serum leptin levels in ob/ob mice transplanted with various volumes of encapsulated control or leptin-producing gut cells 72 Figure 17 The effect of leptin cell therapy on serum insulin levels in ob/ob mice transplanted with various volumes of encapsulated control or leptin-producing gut cells 73 Figure 18 The effect of leptin cell therapy on body weight in ob/ob mice 75 Figure 19 The effect of leptin cell therapy with different doses of RU486 on (A) body weight, (B) food intake and (C) blood glucose in 4-5 week old ob/ob mice 78 Figure 20 Leptin levels measured in 4-5 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of RU486 81 V I I I Figure 21 Insulin levels measured in 4-5 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of RU486 82 Figure 22 The effect of leptin cell therapy with different doses of RU486 on (A) body weight, (B) food intake and (C) blood glucose in 10 week old ob/ob mice 86 Figure 23 Leptin levels from 10 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of RU486 89 Figure 24 Insulin levels from 10 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of RU486 90 Figure 25 Triglyceride levels from 10 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of RU486 91 Figure 26 The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose on ob/ob mice implanted with 3 RU486 pellets 95 Figure 27 The effect of leptin cell therapy on leptin levels in ob/ob mice implanted with 3 RU486 pellets 97 Figure 28 The effect of leptin cell therapy on triglyceride levels in ob/ob mice implanted with 3 RU486 pellets 98 Figure 29 Evaluation of leptin release from encapsulated leptin-producing gut cells maintained in culture for 5 weeks 100 Figure 30 The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose in C57BL/6J mice placed on a high fat diet (HFD) 103 Figure 31 The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose in obese C57BL/6J mice on a high fat diet (HFD) 106 Figure 32 The effect of leptin cell therapy on leptin levels in obese C57BL /6J mice on a high fat diet (HFD) 108 Figure 33 The effect of leptin cell therapy on insulin levels in obese C57BL /6J mice on a high fat diet (HFD) 109 Figure 34 The effect of leptin cell therapy on triglyceride levels in obese C57BL /6J mice on a high fat diet (HFD) 110 Figure 35 The effect of transplanting 18-million leptin-producing gut cells into the IP cavity of obese C57BL/6J mice (pilot study) 113 Figure 36 The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose after transplanting 18-million control or leptin-producing gut cells 115 Figure 37 The effect of leptin cell therapy on leptin levels in obese C57BL /6J mice after transplanting 18-million control or leptin-producing gut cells 118 Figure 38 The effect of leptin cell therapy on insulin levels in obese C57BL/6J mice on a high fat diet (HFD), transplanted with 18-million control or leptin-producing gut cells 119 ix Figure 39 The effect of leptin cell therapy on triglyceride levels in obese C57BL /6J mice on a high fat diet (HFD), transplanted with 18-million control or leptin-producing gut cells 120 Figure 40 Effect of transplanting 18-million encapslated control or leptin-producing gut cells, induced with RU486 24 h prior to transplantation, on (A) body weight, (B) food intake and (C) blood glucose, in obese C57BL/6J mice (pilot study) 122 X LIST OF TABLES Table I Features of the pSwitch plasmid (obtained from the GeneSwitch™ Manual (97)) 22 Table II Features of the pGemB/hLeptin.A/5-His plasmid (obtained from the GeneSwitch™ Manual (97)) 24 Table III Kidney capsule transplant studies in ob/ob mice (studies 11.1 andl l .2) 32 Table IV Encapsulation transplant studies in ob/ob mice (studies 111.1 to III.7) 32 Table V Encapsulation transplant studies in high fat fed C57BL/6J mice (studies IV.1 to IV.5) 33 Table VI Triglyceride levels measured from mice in experiment IV.2 at various weeks throughout the study 111 xi LIST OF ABBREVIATIONS Abbreviation Description AgRP Agouti-related protein ct-MSH a-melanocortin stimulating hormone BMI body mass index CART Cocaine- and amphetamine-regulated transcript C a 2 + Calcium Cs Cyclopsorine DEPC diethylpyrocarbonate DMH dorsal medial hypothalamus ELISA enzyme linked immunosorbent assay F B S fetal bovine serum GIP glucose-dependent insulinotropic polypeptide HFD high fat diet H G D M E M high glucose Dulbecco's modified Eagle's medium i.v. intra-vasucular i.p. intraperitoneal J A K janus kinase LFD low fat diet LH lateral hypothalamus M A P K mitogen-activated protein kinase N P Y neuropeptide Y ObR leptin receptor P B S phosphate-buffered saline PI3K phoshpatidylinositol-3-kinase RIA radioimmunoassay STAT signal transducers and activators of transcription T 2 D type 2 diabetes T G triglyceride V M H ventral medial hypothalamus X I I ACKNOWLEDGEMENTS First, I would like to thank my supervisor, Tim Kieffer, for all of his advice, support, patience and guidance over the past two and a half to three years. It would have been impossible for me to have achieved what I did within these timeframes without Tim's input. I would particularly like to thank Tim for all of his help throughout my writing process - all of the time and effort he spent proof-reading and advising me was greatly appreciated. I could not write an acknowledgements page without thanking certain faculty members that made my time at U B C extremely memorable. Ray Pederson is an individual whom I looked up to for advice and support during my time as a graduate student. Ray really promoted a sense of community and made me feel comfortable and welcome in the department from day one. Ray would always be full of encouraging remarks that really helped get me through some of the more trying times. Of course, I must thank Ray for hosting many fun and memorable parties at his home. Thanks Ray, you really made a difference! Carol Ann Courneya was another constant source of support who was always there to give advice and was always up for a run in Pacific Spirit Park. I am grateful for the opportunities I had to work with "CA" as a teacher's assistant and learn from her skillful and energetic style of teaching. Also, I must thank C A for first introducing me to the great comedian Eric Accilli - thanks for all of the good times you guys. I would like to thank previous members of the "Kieffer Lab" for their contribution to some of the experiments that laid the foundation for this thesis. A huge thank-you to the current members of the "Kieffer Lab": Travis Weber, Suraj Uniappan, Yukihiro Fujita, Scott Covey, Rhonda Wideman, Corinna Lee, Maddy (Superstar!) Speck, Virginia Gunn and Ada Lam. Thanks to you all for your support, advice, help and good times!! I think I'd still be struggling with my Northern blots if it wasn't for Suraj - I hope that one day I will be able to "give back" all of what Suraj has given me in the past year that I've known him. Thanks to Ni Lam, also, for being a great senior grad student "mentor"!! Travis, what can I say? You are the most amazingly organized lab manager who was always available to talk to and provide help when help was needed. There were many times where Rhonda and I would say "Thank heavens for T-Web!!". I'll miss going "head-to-head" with you in Dance!Dance!Revolution! Rhonda, where would I be without you these past few years? We made it through caffeine highs and experimental trials and tribulations. Thanks for all of the coffee talks, laughs and long hours in the lab on Halloween nights!! Good luck to you as you work your way up! I am always and forever grateful to my parents, Joan and Farook, for their constant love, support and encouragement. Mom and Dad, thanks for everything you've given me that has allowed me to get to where I am today. Finally, my absolute deepest thanks must go to Brad. I'm not sure these words will even justify the gratitude that I have for all of the long hours that Brad also devoted to my journey through graduate school. Thanks for being my "sounding board", making me laugh and keeping the stress levels at bay. I know what you put on hold to allow me to pursue my Masters degree and I thank you immensely for that; therefore, I would like to dedicate this thesis to you, Brad. 1 INTRODUCTION Obesity and Type 2 Diabetes Obesity is a chronic metabolic disorder that has been classified as an epidemic due to its accelerated growth throughout the world (1). Obesity has been defined as a state of excessive body weight, from a disproportionate accumulation of fat in the body, at such a high magnitude to promote adverse health consequences (2). Obesity and overweight are defined using the body mass index (BMI; weight/(height)2 in kg/m 2), where overweight individuals have a BMI>25kg/m 2 and obese individuals have a BMI>30kg/m 2. Obesity is widely prevalent in adults and it is increasingly affecting children and adolescents. In Canada, 57% of adult men and 35% of adult women are overweight or obese (3, 4). In 1996, 35.4% of Canadian boys and 29.2% of Canadian girls that were school-aged were considered overweight or obese, an increase from 15% in both boys and girls in 1981 (5). Metabolic changes associated with excess body weight result in an increased risk for developing other diseases such as gall bladder disease, cardiovascular disease, hypertension, some forms of cancer and type 2 diabetes mellitus (T2D) (6, 7). The incidence of T2D is paralleling that of obesity. T2D is a metabolic disorder characterized by hyperglycemia where pancreatic p-cells are unable to produce adequate amounts of insulin within proper time frames required for maintaining normal glucose homeostasis (8, 9). The most prevalent form of diabetes is T2D and 80% of these individuals are obese (10, 11). Diabetes affects over 2 million people in Canada and 17 million people in the United States (11,12). It is estimated that 1 million people over the age of 20 years are newly diagnosed with diabetes in the United States per year (11, 13). The World Health Organization (WHO) has projected that by the year 2025 the number of diabetic individuals will double to almost 300 million worldwide. The 2 W H O also projects that 70% of deaths will be due to the complications associated with diabetes in developing countries by the year 2020 (1). Alarmingly, there has been an increased incidence of diabetes associated with obesity among adolescents worldwide (1, 4). In 1998, diabetes and its secondary complications cost the Canadian health care system between $5.9 and $6.5 billion dollars (14). These figures are expected to dramatically increase as the incidences of obesity and diabetes increase in our population, creating a large financial burden on our Canadian healthcare system. It's not surprising that obesity plays a significant role in the development of T2D (6, 7). A body weight gain of 20 kg increases the risk of developing diabetes by 15-fold (15). Alternatively, a weight loss of 5-11 kg is associated with a 50% decrease in the relative risk of developing T2D (15). Obesity is believed to create an insulin-resistant state where the effects of insulin on glucose uptake, metabolism and storage are impaired, thereby leading to a hyperglycemia (16). Increased body fat is accompanied by an increased release of free fatty acids into the circulation (15). Fatty acids are deposited into non-adipose tissue, such as muscle, heart, and pancreatic B-cells, at the same time that insulin-stimulated glucose uptake into muscle and fat is decreased (17). This fatty acid overload into non-adipose tissue can lead to insulin resistance. Not only is insulin resistance largely linked to the development of T2D it has been linked to the development of other metabolic disorders including hypertension, atherosclerosis and hyperlipidemia (15). It has been well documented that obesity and T2D can be largely prevented through diet restriction or modification and increased exercise and although the general public in today's society are aware of the benefits of such lifestyle changes, the incidences of these 2 chronic disorders continues to escalate. Both genetic and environmental factors influence the development of obesity and diabetes (18,19). The 3 lipostasis theory suggests that every individual has a "set-point" or physiologically preferred level of body weight (20). Body weight appears to remain stable in lean and obese individuals and although modifications in caloric intake can result in weight loss, more than 90% of individuals who lose weight only by dieting eventually return to their original body weight (21). An individual's genes play a key role in the predisposition to attain their ideal body weight. Data collected from twin studies, adoption studies and genetic analyses of certain populations susceptible to obesity, all support the hypothesis that obesity results from genetic factors (22-25). Individuals of various ethnicities that have become "westernized" have a high propensity to developing obesity and diabetes (26, 27). The increasing prevalence in obesity, however, has not been due to any changes in the genetics of our population, but rather due to changes in our environment, specifically the abundance of food and lack of physical activity (2). It is likely that the propensity for obesity has always been a genetic trait in humans; a positive trait selected throughout evolution to give humans the ability to store fat in times of nutritional abundance (2). Genetics may lay the foundation for susceptibility to obesity and diabetes but it's the interactions of these genes with environmental factors that determine whether an individual will develop obesity and/or diabetes. Parabiosis experiments and the sateity signal In 1953, Kennedy proposed the lipostasis theory which describes a negative feedback control system; this system involves a peripheral signal, which is generated in proportion to the amount of fat stores in the body, that acts on the brain to influence food intake and energy expenditure (20, 28, 29). In other words, energy expenditure and energy intake are modulated continuously to achieve a constant adipose tissue mass (30). Experiments were undertaken by several researchers to further elucidate 4 the mechanisms involved in body weight regulation, and to hunt for the unknown peripheral signal; these experiments required animal models of obesity and diabetes. Ob/ob and db/db mice are obese and diabetic exhibiting hyperphagia, hyperglycemia, insulin resistance and an increase in adiposity. In the 1970's, Coleman et al. performed cross-circulation, or parabiosis, experiments on these mouse models. In one study, Coleman et al. partially fused the circulations of the ob/ob mice to wild-type mice and observed reduced feeding and weight loss in ob/ob mice (31, 32). In another study, Coleman etal. partially fused the circulations of ob/ob mice and wild-type mice with db/db mice. Coleman et al. discovered that the ob/ob and wild-type mice had a decrease in appetite, an increase in energy expenditure and a decrease in body fat mass but the db/db mice did not respond with any of these changes (33, 34). It was postulated that ob/ob mice were responding to an unknown circulating satiety factor that was absent in their own circulation but was produced in wild-type and db/db mice. Further, it was hypothesized that db/db mice were resistant to this same unknown appetite-suppressing hormone (31, 32). The exact defect and site of synthesis of this hormone was unknown and would remain unknown for several years. Leptin In 1994, several decades after the parabiosis experiments were performed, Friedman and his colleagues discovered the hormone leptin through positional cloning techniques (19). They found that a mutation in the obese (ob) gene in ob/ob mice prevented the production and secretion of the full, active form of the leptin protein, which led to the obesity and diabetes phenotype in these mice. Friedman et al. found that leptin was secreted in the periphery, primarily from white adipose tissue. The discovery of this peripherally secreted protein supported findings from the parabiosis 5 experiments done by Coleman et al. that suggested that ob/ob mice were deficient in a circulating satiety factor that db/db mice produced but were resistant to (34). In 1995, researchers treated leptin-deficient ob/ob mice with leptin and observed dramatic weight loss and normalization of blood glucose levels in these mice (35-38). However, when leptin was injected into db/db mice, they did not respond with reductions in body weight or blood glucose levels. The diabetes (db ) gene was located in the feeding centres of the hypothalamus (18, 39). Again, these data confirmed Coleman et al.'s predictions: db/db mice produce high amounts of a circulating satiety signal (leptin) but are resistant to it. The ob gene yields a 3.5 kb mRNA and encodes a 167 amino-acid protein termed leptin. Human and mouse ob genes are localized to chromosomes 7q31.3 and 6, respectively (40, 41). Leptin is a globular protein similar to members of the long chain helical cytokine family and is comprised of 4 a-hel ices and 2 short p-sheets bound by a disulfide bond (29). Leptin is produced and secreted primarily from adipose tissue as a 16 kD protein (19). Leptin expression has also been detected in placenta (42), stomach (43), muscle (44) and brain (40, 45), but its role in these peripheral t issues is not fully understood. Leptin circulates either as an active, free form, or as an inactive form bound to leptin-binding proteins (18). Human leptin is highly homologous to mouse leptin (84% homology) and rat leptin (83% homology) (19, 46). Leptin's primary role is believed to be the relaying of information about the amount of energy stored in adipose tissue to regions in the brain (29, 47, 48). The leptin signal plays a role in the regulation of feeding behaviour through a distinctive feedback loop. P lasma leptin levels correlate closely with adipose tissue mass so as adipose stores increase, leptin is produced and secreted in greater quantities (7, 18, 33, 49). It has also been found that adipocyte size is important in regulating leptin secretion since larger 6 adipocytes produce more leptin than smaller adipocytes in the same individual (49, 50) Generally, in humans, circulating leptin concentrations remain at a steady basal level within an individual but differ from individual to individual depending on their body mass index (BMI). Males and females with a BMI between 18 and 25 kg/m 2 were found to have approximately 3.84±1.79 ug/L and 7.36±3.73 ug/L of circulating leptin respectively (51). Obese males and females with a BMI>27 kg/m 2 had average circulating leptin levels of 31.3±24.1 ng/mL (47). Generally, when adipose tissue mass increases, the amount of circulating leptin also increases. Bado ef al. measured the concentration of leptin in the stomach of rats after a meal and they found that gastric leptin concentrations decreased by approximately 66% from baseline measurements within the first 15 minutes of eating. Concurrently, they measured approximately a 70% increase in plasma leptin concentrations suggesting that leptin release from the stomach directly affects plasma leptin concentrations (43). In rodents and humans, rapid decreases in leptin expression occur within several hours in response to fasting and disproportionately to corresponding alterations in body weight and fat mass (48, 52-55). Since rapid decreases in leptin levels initiate food-seeking and energy-conserving behaviour, leptin is believed to be an important survival signal during periods of food deprivation (56). Female mice stop ovulating when in a starved state yet starved female mice, when administered leptin, will continue to ovulate even though their body weight continues to decrease. Again, these data suggest that leptin's primary role is in times of food deprivation (56). Decreased leptin levels, therefore, may play a more crucial physiological role than increased leptin levels to ensure sufficient maintenance of caloric reserves. 7 Leptin receptor Tartaglia etal. first isolated the leptin receptor (ObR), in 1995, by an expression cloning strategy using mouse choroid plexus (57). Through positional cloning, Lee et al. revealed that the leptin receptor gene (db) encodes at least six alternatively spliced forms of mRNAs : ObRa , ObRb, ObRc, ObRd, ObRe and ObRf (41). Of the six identified, the long form of the receptor, ObRb, is the most extensively characterized and has been found to be essential for the weight-reducing effects of leptin. The ObRb is expressed at high levels in the hypothalamus and, to a lesser degree, in peripheral t issues (58). Specifically, ObRb receptors in the hypothalamus are located within regions that regulate food intake and energy expenditure, namely the arcuate nucleus, the ventromedial hypothalamic nucleus (VMH), the dorsal medial hypothalamic nucleus (DMH), and the lateral hypothalamus (LH) (7, 59, 60). The db/db mouse and fa/fa rat, which have a similar phenotype to the obese and diabetic ob/ob mouse, do not express the functional ObRb receptor as a result of mutations in the db gene, but they express the short form O b R a receptor. This indicates that the long form ObRb receptor is critical for leptin action and explains why these animals do not respond to leptin administration (41, 61). The other isoforms of ObR are also widely distributed throughout the periphery but their function remains unclear (33, 62). The ObRb has been identified as a member of the class I cytokine family of receptors and is involved in signalling via the janus-kinase/signal transducer and activator of transcription 3-kinase (JAK/STAT-3) pathway (57). Leptin binds to a homodimer of ObRb (63, 64), leading to the phosphorylation of J A K 2 . Activation of J A K 2 initiates the phosphorylation of the intracellular domain of the receptor thereby allowing the binding and activation of the STAT proteins. The STAT proteins dimerize and translocate to the nucleus where they regulate gene transcription (65). Activation of 8 J A K 2 also promotes phosphorylation of phosphoinositide 3-kinase (PI3-kinase) and mitogen-activated protein kinase (MAPK) pathways (66, 67). Leptin receptors within the LH and V M H are involved in balancing food intake and energy expenditure through modulating activity of the parasympathetic and sympathetic nervous systems. These hypothalamic nuclei express various neuropeptides and neurotransmitters that regulate food intake and energy expenditure. Leptin enters the brain through a saturable transport mechanism at the blood-brain barrier (68). When leptin levels are low, orexigenic peptides such as agouti-related protein (AgRP) and neuropeptide Y (NPY) are stimulated, while anorectic peptides such as a - M S H (a melanocortin receptor agonist) and cocaine- and amphetamine-regulated transcript (CART) are inhibited, thereby leading to an increase in food intake and a decrease in energy expenditure (7, 59, 60, 69). If leptin levels are high, as in an overfed state, leptin's overall effect is to decrease food intake and increase energy expenditure through the actions of the above pathways (52, 70). These changes that occur in response to overfeeding or refeeding following a period of starvation, are observed after several days (71, 72). Through these hypothalamic pathways, leptin is able to communicate the state of energy stores from the adipocyte to the brain, causing appropriate changes in food intake and energy expenditure to occur. Many researchers propose that, historically, the actions of leptin were most important in protecting fuel stores during periods of food deprivation and that leptin's main purpose was to ensure the maintenance of sufficient energy stores (56). Leptin in the treatment of obesity and diabetes Ob/ob and db/db mice are phenotypically characterized by obesity, hyperglycemia, hyperinsulinemia and insulin resistance, which closely resembles 9 obesity and diabetes in humans (34). One year following the discovery of leptin, several research groups observed that injecting leptin into ob/ob mice resulted in reductions in body weight and food intake as well as normalization of hyperglycemia and hyperinsulinemia (35-38). This leptin-induced weight loss was specific for reductions in adipose tissue mass (36, 73, 74), and was greater than in control ob/ob mice pair-fed the same amount of food consumed by the leptin-treated ob/ob mice (74). This observation is attributable to the ability of leptin to induce weight loss by increasing energy expenditure in addition to reducing food intake. Pelleymounter et al. demonstrated that treating ob/ob mice with low doses (0.1 mg/kg) of leptin by injection did not prevent body weight gain but did cause a drastic reduction in plasma glucose levels (by 66%) and plasma insulin levels (by 41%) (37). Normalization of blood glucose levels also occurred prior to a decrease in body weight in ob/ob mice administered leptin by gene therapy (75, 76). Dramatic decreases in plasma glucose and insulin levels are even observed within 24 h, with no changes in body weight (77, 78). The data from all of these studies suggest that leptin may have a direct effect on glucose homeostasis independent of its effects on body weight. Leptin gene therapy has been shown to be more efficacious in the treatment of obesity and diabetes in ob/ob mice compared to treatment by leptin injection. A study, performed by Morsy et al., compared the efficacy of treating ob/ob mice with recombinant leptin protein delivered by injections versus by gene delivery (76). Greater weight loss was achieved by gene delivery (~11 g loss) over a 14-day timeframe compared with daily leptin injections (~5 g loss). This accelerated weight loss associated with gene delivery was attributed to the sustained expression and circulation of the gene-delivered leptin. This study also revealed that ten times less leptin was required to cause this increased rate of weight loss with leptin gene therapy than with 10 injection of leptin. Treatment by leptin injection requires large doses to be administered daily but since its half life has been reported to be approximately 3 hours, it is rapidly degraded in the bloodstream, permitting only a short duration of leptin activity (56). Gene therapy is likely more efficacious because it allows continuous low-dose circulation of leptin which parallels normal physiological kinetics much more closely than by bolus injection. Human congenital leptin deficiency, a rare disorder, is a clinical phenotype that is very similar to that seen in ob/ob mice (70, 79, 80). Leptin-deficient humans suffer from early-onset obesity, increased appetite, impairment in sexual development, defects in immune-mediated responses, hyperinsulinemia and deregulation of the hypothalamo-pituitary axis, all of which are observed in ob/ob mice (52, 70, 80, 81). Individuals that were leptin-deficient and treated with leptin by injection responded with reductions in food intake and sustained body weight loss, primarily loss of fat mass (70). No significant effects on basal metabolic rate or energy expenditure have been observed (52, 70). All other phenotypic abnormalities were improved with leptin injection therapy (70, 79, 80). These findings provide evidence that leptin injection therapy is beneficial in treating the multiple phenotypic deviations associated with congenital human leptin deficiency. Unlike individuals suffering from congenital leptin deficiency, the majority of humans suffering from obesity have high levels of circulating leptin (82). A s fat mass increases and leptin levels rise in obese individuals, ieptin's ability to suppress food intake and prevent obesity wanes due to leptin resistance. The physiological basis of leptin resistance is currently unknown but is likely the result of many different factors. It has been suggested that entry of leptin into the cerebrospinal fluid (CSF) may be limited, leading to leptin resistance (83, 84). Furthermore, the effective transport of 11 leptin across the blood-brain barrier has been questioned in pre-obese and obese animals (68, 85, 86). Leptin signal transduction via its J A K / S T A T pathway could also be defective at one or many different levels, thereby leading to a state of leptin resistance (66). C57BL /6J mice develop obesity and diabetes when fed a high-fat diet (54, 55, 87, 88). Apparent resistance to endogenous and exogenous leptin can be demonstrated in mice placed on a high fat diet for several weeks (87, 88). When individuals suffering from obesity were injected with leptin, only some of the subjects responded with decreases in body weight. These decreases were achieved with the highest dose of administered leptin and the data were inconsistent since not all subjects treated with this doses of leptin lost weight (53). The highest dose of exogenous leptin administered to the individuals in this study done by Heymsfield et al. was 0.30 mg/kg/day, which equated to 115.5 ng/mL of serum leptin concentration (measured at week 24 of the study, 4 h after injection; (53)). In comparison, leptin-deficient obese humans lost body weight with daily leptin injections of only 0.01-0.04 mg/kg/day, which illustrates that obese individuals in the Heymsfield study (53) experienced leptin resistance. A s previously reported, lean humans have leptin levels between 3 and 7 pg/L (51); so even though obese individuals in the Heymsfield study had over 16-times more circulating leptin in their system than lean individuals, they continued to have a much higher body mass compared to the lean individuals. These data suggested that administration of exogenous leptin may cause weight loss in some obese individuals but not in others, indicative that some obese individuals may be resistant to the effects of leptin. Leptin cell therapy Treatments for peptide hormone deficiencies typically involve frequent administration of the therapeutic protein by injection. Leptin has been effectively 12 administered by injection in leptin deficient animals and humans but requires repetitive dosing and does not accurately mimic leptin's normal physiological release profile (36, 37, 76). Leptin injection causes transient increases in plasma leptin levels and is ineffective in leptin-resistant states. Leptin administration by gene therapy allowed continuous secretion of low levels of leptin which has been shown to be more effective in treating obesity and diabetes in ob/ob mice than repetitive leptin injections (75, 76). It is possible that continuous delivery within the body may be a strategy to overcome leptin resistance associated with obesity. The main limitation of gene therapy is in delivering therapeutic genes to patients in a safe and effective manner (89). Moreover, if a therapeutic protein causes secondary deleterious effects, it may be difficult and sometimes impossible to reverse these changes. In considering the actions of leptin, if too much leptin is produced it can lead to immune and reproductive dysfunction, bone mass loss, 6-cell dysfunction and even leptin resistance (89, 90). Cell therapy utilizes living cells to deliver therapeutic proteins. Some cells are ideal vehicles to administer therapeutic proteins because they can produce and secrete protein products and respond to physiological changes. In most immortalized cell lines, however, physiologically regulated processes are often not operative (91). Regulation of expression of transferred genes has been closely studied in the field of cell therapy throughout the past decade (92). Systems have been developed in an attempt to regulate the synthesis and secretion of protein products. In developing a leptin cell therapy system, it is desirable to incorporate an "on/off system in order to adapt to metabolic fluctuations that can frequently occur in individuals. It is possible to genetically engineer cells to respond to a regulatable external cue, such as a drug, thereby switching gene expression "on" or "off whenever necessary, or even maintaining expression at a constant level (93). Regulatable gene expression systems 13 can allow dose-dependent release of the product of the gene of interest and specifically meet the needs of each individual. Several groups have designed systems where administration of a drug is used to stimulate the activity of a transcription factor, which in turn will stimulate the production of the therapeutic protein of interest (92-95). These systems allow precise dosages of gene products to be administered (94, 95). The GeneSwitch System™ is a regulatory system for gene transfer that was developed by Wang et al. in 1994 (94). This inducible system consists of a regulator component and a target component (96). When an exogenous stimulus such as RU486 is administered, it binds to the regulator component of the system thereby activating the expression of the target gene (96, 97). Administering RU486 at different doses or for different durations modulates the level and duration of target gene expression (96). This system also allows for the 'switching on' or 'switching off of target genes by either administering or discontinuing RU486. In designing a cell therapy it is important to consider and choose an appropriate cell-type for the delivery of the therapeutic product. Intestinal endocrine cells are capable of producing, storing and secreting proteins in response to external stimuli. Intestinal K-cells, for example, secrete glucose-dependent insulinotropic polypeptide (GIP) in response to in the ingestion of nutrients. Previously, it has been shown that a GIP-expressing intestinal tumor-derived K-cell line could be engineered to produce another protein, insulin (98). This K-type tumor cell line was initially derived from the murine intestinal cell line STC-1 (secretin-producing tumor cells), a mixed population of intestinal endocrine cells (99). STC-1 cells were transfected with an expression plasmid containing a portion of the rat GIP promoter fused to the gene encoding the enhanced green fluorescent protein (EGFP) ; K-type tumor cells expressing large amounts of GIP m R N A were thus visually identified as GIP tumor cells, GTC-1 cells, and it was these 14 GTC-1 cells that were engineered to produce insulin (98). Cheung et al. observed that when these genetically modified K-type tumor cells were induced with glucose, insulin was produced and secreted in a glucose-dependent manner (98). Interestingly, in 1998 Bado et al. discovered that in addition to fat, stomach cells also produce leptin, evidently in response to a meal (43). Considering that intestinal K-type tumor cells are meal-responsive, it could be possible and appropriate to genetically engineer these cells to produce leptin. Cellular release of leptin may also provide a more constant release of leptin compared to injections, similar to the physiological steady-state levels previously observed (29, 100). Genetically engineering intestinal K-type tumor cells to secrete leptin could prove to be an effective means of treating obesity and diabetes in leptin-deficient or leptin-resistant animals by boosting both basal and post-prandial levels of leptin expression and release. Transplantation strategies There are several different strategies that could be used in transplanting cells for cell therapies. One such transplantation strategy is the kidney capsule transplantation technique. The kidney capsule provides a very effective barrier in which to transplant cells. The kidney capsule is the outer layer that surrounds the highly vascularized kidney. Transplanting cells into this capsular region is advantageous since the capsule itself acts as a barrier to the immune system thereby offering the cells some protection from destruction by the immune system. The other advantage to transplanting cells in the kidney capsule is that it is a highly vascularized region thereby promoting angiogenesis within the transplanted cells, again encouraging their survival (101). Cell encapsulation is another technique of protecting cells prior to transplantation. Encapsulation allows cells to be immuno-isolated within a 15 semipermeable membrane that allows influx of some nutrients and outward release of secreted proteins (91). Encapsulating cells prior to transplantation is advantageous because there is no longer a need for immunosuppression. Several research groups have previously encapsulated pancreatic islet tissue in hydrogels and have observed that this tissue remains functional between 5 and 20 weeks (63, 102). The limited graft survival was attributed to hypoxia rather than by immune cell destruction. Encapsulated cells have been observed to survive between 6 weeks (103) and 6 months (91). Microencapsulated rat islets were transplanted into the intraperitoneal cavity of mice and after 9 weeks the encapsulated islets were retrieved. Glucose-stimulated insulin release was still observed but the output was less than half of the release observed in the pre-treatment control islets (104). Others have shown that insulin release from pancreatic islets was the same when comparing release from free versus microencapsulated islets (103). Microencapsulated adult pig islets transplanted into diabetic mice survived up to 190 days providing evidence that encapsulation can prevent rejection of donor tissue by host immune destruction (105). Other investigators have shown that by optimizing the encapsulation process, survival of transplanted cells may continue to improve (106) thereby providing longer-term therapies in the treatment of such diseases as obesity and diabetes. Moreover, the issue of protecting transplanted cells from host immune destruction may not be an issue if techniques could be developed for individuals to donate their own cells that could be modified to produce therapeutic protein products prior to returning the cells back to the same individual. 16 THESIS INVESTIGATION The prevalence of obesity and T2D is increasing throughout the world. 80% of individuals suffering from T2D are obese and it has been recognized that there is a strong association of obesity to diabetes. However, the molecular basis for the link between these 2 chronic disorders remains a mystery. Leptin has been suggested as one candidate that bridges the gap between obesity and T2D. Leptin has been shown to reverse the obesity and diabetes phenotype in rodent models of congenital leptin deficiency (35-38). Although leptin injection therapy has promoted weight loss and normoglycemia in leptin-deficient rodents and humans, leptin injection therapy has only been moderately successful in treating leptin-resistant obese individuals. The research in this thesis examines the possibility that a leptin cell therapy may be efficacious in treating leptin-deficient as well as leptin-resistant rodent models of obesity and diabetes. The research in this thesis examines the ability of genetically engineered leptin-producing gut cells to produce leptin in the presence of an exogenous drug and the therapeutic effect of leptin cell therapy in mouse models of obesity and diabetes. First, experiments were done to characterize genetically engineered leptin-producing gut-cells. Second, a comparison of the effect of transplanting leptin-producing gut cells into the kidney capsule of ob/ob mice that did not receive immunosuppressant therapy with mice that did receive immunosuppressant therapy was done. Third, the effect of encapsulating leptin-producing gut cells prior to intraperitoneal transplantation into ob/ob mice was explored; within these experiments further evaluation was done to determine whether the studied parameters were RU486 dose-dependent in vivo, and to determine how long the encapsulated leptin-producing gut cells released leptin into these ob/ob mice. Finally, the effect of transplanting encapsulated leptin-producing gut cells into the intraperitoneal cavity of high-fat-fed, obese C57BL6/J mice was evaluated. 17 MATERIALS AND METHODS The GeneSwitch™ System The GeneSwitch System is a regulatory system for gene transfer that was developed by Wang et al. in 1994 (94). This inducible system consists of a regulator component and a target component (96). The regulator, which can be expressed in any cell by using appropriate tissue-specific promoters, contains a Gal4 DNA binding domain, an activation domain and a truncated human progesterone receptor. The activated regulator was designed to bind to and only activate target genes with a Gal4 DNA binding site, only in the presence of Mifepristone (RU486). The target can be any gene under the control of a minimal promoter (TATA box) and four high-affinity Gal4 D N A binding sites. When RU486 is administered, it binds to the progesterone receptor on the regulator; the regulator becomes activated after undergoing a conformational change and it binds to the Gal4 DNA binding sites upstream of the target gene, which is then expressed (96, 97) (Figure 1). 18 Human leptin hllp://w vvvv. gene sw i t c h . c o m / a u t o / i n d c x . b t m i l GAL4-DBD/hPR/p65 -AD fusion gene Figure 1. Schematic representation of the GeneSwitch"" System transfected into GTC-1 cells (A). Plasmid 1 contains the pSwitch vector that was transfected into GTC-1 cells resulting in the GTC-1 pSwitch cell line (control gut cell line, B). Plasmid 2 contains the human leptin DNA vector that was transfected into the control gut cell line to produce the GTC-1 pSwitchGIProhLeptin cell line (leptin-producing gut cell line, C). 19 Generation of the control gut cell line and the leptin-producing gut cell line using the GeneSwitch™ System GIP-expressing GTC-1 cells, derived from the STC-1 cell line (98, 107), were transfected with a modified regulatory GeneSwitch™ System (Invitrogen Canada Inc., Ontario, Canada) to create two different cell lines: GTC-1 pSwitchGIPro cells (control gut cell line) and GTC-1 pSwitchGIProhLeptin cells (leptin-producing gut cell line). Control gut cells (GTC-1 pSwitchGIPro) were generated by replacing the constitutively active C M V promoter with a rat GIP promoter sequence (2.6 kb; generously provided by Dr. Boylan, (107)) into the pSwitch vector (GeneSwtich™ system) at Sbfl and Pac I restriction enzyme sites to create the pSwitchGIPro plasmid (Figure 2A; Table I). The pSwitch vector originally contained a GAL4 Upstream Activating Sequence which was also removed when the rat GIP promoter was inserted. This pSwitchGIPro plasmid, containing a Hygromycin-resistance gene, was stably transfected into GTC-1 cells using L I P O F E C T A M I N E PLUS™ reagent (Invitrogen Canada Inc.). The newly constructed GTC-1 pSwitchGIPro (control) gut cells were selected with Hygromycin B (500 pg/mL; Invitrogen Canada Inc.). Leptin-producing gut cells (GTC-1 pSwitchGIProhLeptin) were constructed through several steps. First, human leptin cDNA (Human Fat Cell QUICK-clone; Clonetech, CA , USA) was replicated by R T - P C R then ligated into the multiple cloning site of the inducible expression cassette (pGeneB/V5His) obtained from the GeneSwitch™ System (Figure 2, B; Table II). Human leptin c D N A was cloned into the pGeneB/V5His expression cassette at Apa I and Spe I restriction enzyme sites. The V5His region of the pGeneB plasmid was removed using Apa I and Pme I restriction enzymes thereby creating the pGeneB/hLeptin/-V5His plasmid. This plasmid contains the zeocin resistance gene. Second, this plasmid was stably transfected into the 20 hygromycin-resistant control gut cell line (GTC-1 pSwitchGIPro cell line), containing the pSwitch plasmid, using LIPOFECTAMINE PLUS™ reagent. The new GTC-1 pSwitchGIProhLeptin cells contained both the pSwitch regulatory plasmid and the modified inducible human leptin plasmid, which will be referred to as the leptin-producing gut cell line. Selection of leptin-producing gut cells was conducted using Hygromycin B (500 pg/mL) and Zeocin (800 pg/mL; Invitrogen Canada Inc.). All cells were cultured at 37 C, 5%C0 2 /95% air in high-glucose Dulbecco's modified Eagle's medium (4.5 g/L D-glucose, HG-DMEM; Invitrogen Canada Inc.) supplemented with 10% fetal bovine serum (FBS; Invitrogen, Canada Inc.), 100 pg/mL streptomycin (Invitrogen Canada Inc.), 100 U/mL penicillin (Invitrogen Canada Inc.) and 3.7 g/Lsodium bicarbonate. 21 Figure 2 A. Schemat ic representation of the steps performed in constructing the pSwitchGIPro plasmid. This first modified plasmid of the GeneSwi tch system was transfected into G T C - 1 cells to produce the pSwitch cell line (control gut cell line). 22 Table I. Features of the pSwitch plasmid (obtained from the GeneSwitch™ Manual M M : , , Feature Benefit Herpes Simplex Virus thymidine kinase (TK) minimal promoter Allows expression of the GAL4-DBD/hPR-LBD/p65-AD fusion gene (McKnight, 1980) Synthetic intron IVS8 Enhances expression of the GAL4-DBD/hPR-LBD/p65-AD fusion gene GAL4-DBD/hPR-LBD/p65-AD fusion gene Encodes a 73 kDa fusion protein containing the yeast GAL4 DNA binding domain (Laughon and Gesteland, 1984; Marmorstein et al., 1992), a truncated human progesterone receptor ligand binding domain (Kastner era/. , 1990; Misrahi et al., 1987; Wang et al., 1994), and the human NF-£eB p65 activation domain (Burcin et al., 1999; Deloukas and Loon, 1993; Ruben et al., 1991) to allow mifepristone-regulated expression of the gene of interest Bovine growth hormone (BGH) polyadenylation sequence Permits efficient polyadenylation of mRNA (Goodwin and Rottman, 1992) B G H reverse priming site Permits sequencing of the non-coding strand f l origin Allows rescue of single-stranded DNA SV40 early promoter and origin Allows efficient, high-level expression of the hygromycin resistance gene in mammalian cells and episomal replication in cells expressing SV40 large T antigen Hygromycin (HPH) resistance gene Permits selection of stable transfectants in mammalian cells (Gritz and Davies, 1983) SV40 early polyadenylation signal Allows polyadenylation of mRNA pUC origin Permits high-copy number replication and maintenance in E. coli bla promoter Allows expression of the ampicillin (bla) resistance gene Ampicillin (bla) resistance gene (£]-lactamase) Allows selection of transformants in E. coli 23 SacD Neo I Bstz 1 Sph I Ant II Ant I GTA ,spe I spe i Not I Ban 41 Icpn 2 Asp78l Hind II ligatcd spe I A pa I cut t v5His Pmel ,apa I vs His ,Pme I Figure 2 B. Schemat ic representation of the steps performed in constructing the pGEMB/hLept in / -V5His plasmid. This second modified plasmid of the GeneSw i t ch system was transfected into the pSwitch cell line (control gut cell line), containing the pSwitch p lasmid, to produce the leptin-producing gut cell line. 24 Table II. Features of the pGemB/hl_eptin/-V5-His plasmid (obtained from the GeneSwitch™ Manual (97)). Feature Benefit GAL4 Upstream Activating Sequences (UAS) Contains six copies of a 17 bp sequence that allows binding and transcriptional activation of the gene of interest by the GAL4 DBD/hPR-LBD/p65 AD fusion protein (Giniger et al., 1985; Wang etal., 1994) Adenovirus E1b TATA sequence Permits transcriptional initiation of the gene of interest (Lillie and Green, 1989) Synthetic intron IVS8 Enhances expression of the gene of interest pGene forward priming site Allows sequencing in the sense orientation Multiple cloning site Allows insertion of your gene of interest V5 epitope (Gly-Lys-Pro-lle-Pro-Asn-Pro-Leu-Leu-Gly-Leu-Asp-Ser-Thr) Allows detection of your recombinant protein with the Anti-V5 antibodies (Southern et al., 1991) C-terminal polyhistidine (6xHis) tag Permits purification of your recombinant protein on metal-chelating resin such as ProBondo or Ni-NTA In addition, the C-terminal polyhistidine tag is the epitope for the Anti-His(C-term) antibodies (Lindner etal., 1997) Bovine growth hormone (BGH) polyadenylation sequence Permits efficient polyadenylation of mRNA (Goodwin and Rottman, 1992) BGH reverse priming site Permits sequencing of the non-coding strand f1 origin Allows rescue of single-stranded DNA SV40 early promoter and origin Allows efficient, high-level expression of the Zeoc in 0 resistance gene in mammalian cells and episomal replication in cells expressing SV40 large T antigen EM7 promoter Synthetic prokaryotic promoter for expression of the Zeocino resistance gene in E. coli Zeoc in 0 resistance (Sh ble) gene Permits selection of stable transfectants in mammalian cells (Mulsant etal., 1988) and transformants in E. coli (Drocourt et al., 1990) SV40 early polyadenylation signal Allows polyadenylation of mRNA pUC origin Permits high-copy number replication and maintenance in E. coli. bla promoter Allows expression of the ampicillin (bla) resistance gene Ampicillin (bla) resistance gene (£]-lactamase) Allows selection of transformants in E. coli 25 In vitro studies Northern blot determination of GIP mRNA expression GIP m R N A expression in GTC-1 cells, control gut cells (GTC-1 pSwitchGIPro cells) and leptin-producing gut cells (GTC-1 pSwitchGIProhLeptin cells) were compared by Northern blot analysis. All cells were grown to approximately 80% confluency. Cel ls were either uninduced or induced with 10' 8 M RU486 (Mifepristone, Invitrogen Canada Inc.) 24 h prior to R N A extraction. Total R N A was extracted using TRIzol Reagent (according to the manufacturer's instructions; Invitrogen Canada Inc.). The integrity and quantity of R N A was measured using a spectrophotometer at 260/280 nm. For Northern blot analysis, 40 pg of total R N A was electrophoresed on a 1 % denaturing agarose gel (5 mL 10x M O P S / E D T A , 37.5 mL DEPC-treated water, 15% formaldehyde, 5 pL ethidium bromide). Confirmation of m R N A band size was made at this time and the denatured R N A was then transferred from the gel onto a nylon membrane ( O S M O N I C S Inc., Westborough, MA). Slot blot analysis was also carried out to further quantify the amount of GIP m R N A expression in these cells. For slot blot analysis, 20 pg of R N A was loaded into the slot blot apparatus and the R N A was transferred onto a nylon membrane by vacuum. All nylon membranes were baked at 80 C in an oven for 2 h and UV cross-linked for 30 s. GIP cDNA (660 bp, a gift from Dr. Michael Boylan, Boston University School of Medicine and Boston Medical Center) and 6-actin c D N A (1076 bp; Ambion, Cambridgeshire, UK) probes were radiolabeled with 3 2 P by the random primer technique using the Rediprime II kit (GE Healthcare, Baie-d'Urfe, P Q , Canada). Nylon membranes were then hybridized first with 3 2 P-label led GIP c D N A probe at 65 C overnight ( 3 2 P purchased from G E Healthcare). Nylon membranes were washed three times with wash solution (1 M N a H P 0 4 , 0.5 M EDTA, 20% S D S ) and then 26 set to autoradiograph for up to 7 days. Northern and slot blot membranes were stripped with stripping solution (20% S D S , 20x S S C , 1 M TRIS/CI, pH 7.5) and re-probed with the 3 2 P- label led B-actin probe. Nylon membranes were again washed with wash solution and autoradiographed for 24 h. Blots were quantified using the ImageQuant program (Molecular Dynamics-Amersham Biosciences). Blots of GIP m R N A were normalized to p-actin m R N A levels to take into account any differences in loading. Effect of RU486 dose and exposure time on leptin mRNA expression in leptin-producing gut cells Dose and time response Northern analyses were done to determine the levels of human leptin m R N A expression in genetically engineered leptin-producing gut cells in the presence or absence of RU486. Cells were grown to 60% confluence on 6 cm plates and, for the dose response study, induced with various doses of RU486 (0 M, 10" 1 2 M, 10" 1 0 M, 10"8 M or 10 ' 6 M); media was collected after 8 h of incubation with RU486 and R N A was extracted from adherent cells, as previously described. For the time course study, cells were also grown to 60% confluence on 6 cm plates and induced with 10" 8 M RU486 with media being collected at 0 h, 4 h, 8 h and 24 h. R N A extraction was carried out as previously described. Fifteen u.g of R N A was loaded onto a 1% denaturing agarose gel And Northern blotting was carried out as previously described, using a 3 2 P-label led human leptin c D N A probe (840 bp human beta-actin cDNA; Clontech). Western blot analysis of leptin protein expression in leptin-producing gut cells Cells were prepared in the same way as for the Northern blot analysis of human leptin m R N A expression. When cells reached 60% confluence, they were either 27 induced with various doses of RU486 (0 M , 10" 1 2 M, 1 f J 1 0 M, l O - 8 M or 10" 6 M) or induced with 10" 8 M RU486 for various time periods (0 h, 4 h, 8 h, or 24 h). Cells were washed 2x with ice cold P B S then lysed with lysis buffer (50 mM H E P E S , 1% Triton-X, 2 mM EDTA, 200 mM NaF, 10 mM N a 2 P 4 0 7 , 1mM P M S F , 1 mM N a 3 V 0 4 ) 10 pg/mL protease inhibitor cocktail; all reagents from Sigma-Aldrich Canada Inc., O N , Canada). Lysates were harvested from the plates, assessed for protein concentration by Bradford protein assay and stored at -80 C until further analysis. Protein samples were run on a 10% S D S P A G E gel and transferred overnight at 4 C to a nitrocellulose membrane (OSMONICS Inc.). The membrane was blocked in TBS-Tween, pH 7.4, (20 mM TRIS, 150 mM NaCI, 0.1% Tween 20) containing 5% non-fat dry milk. Immunoblotting was performed using an affinity-purified leptin antibody (1B:leptin, A-20; Santa Cruz Biotechnology, Santa Cruz, C A , USA) , and a HRP-conjugated anti-rabbit IgG antibody (Santa Cruz Biotechnology). Protein was visualized by enhanced chemiluminescence (ECL western blotting system; Amersham Pharmacia Biotech, Inc., Uppsala, Sweden). The chemiluminescent signal was captured on Kodak BioMax M R film (Eastman Kodak Co. , Rochester, NY). ELISA analysis of leptin release from leptin-producing gut cells Cells were grown in a T-25 flask to approximately 70% confluence and were then seeded onto 24-well plates. Cells were grown for another 48 h prior to induction with either various doses of RU486 or with 10"8 M RU486 for various time periods, as previously described. Media (100 pL) was harvested from each well and these media samples were used to assay for human leptin using a human leptin EL ISA kit ( A L P C O Diagnostics, Windham, USA) . This ELISA kit has 100% cross-reactivity with human 28 leptin and <0.2% cross-reactivity with mouse and rat leptin. The lowest detectable level of leptin in this kit is 0.2 ng/mL. Encapsulation of leptin-producing or control gut cells Cells were grown to 80% confluence and were trypsinized using 0.25% trypsin/1 mM EDTA. The cells were pelleted by centrifugation and the supernatant aspirated and discarded. The cell pellet was resuspended in M O P S wash buffer (10 mM M O P S + 0.85% NaCI). The MOPS-ce l l suspension was transferred into 10 mL of 1.5% sodium alginate solution (1.5% low viscosity alginate in M O P S washing-buffer, pH 7; Inotech Biosystems Inc., MD, USA) and then transferred to a 20 cc syringe. The syringe containing the polymer-cell suspension was then attached to the syringe pump, which forced the mixture through the pulsation chamber on the Inotech encapsulator (Research IE-50R; Inotech Biosystems Inc.). The polymer-cell suspension passed through the bead-forming nozzle on the encapsulator causing the formation of equal sized droplets or beads. Prior to exiting the nozzle, the beads passed through an electrical field causing them to have a surface charge. The electrostatic charge on the beads provided repulsive forces that caused the beads to disperse as they dropped into the encapsulator chamber containing polymerization solution (10 mM M O P S , 100 mM CaCI 2 ; Inotech Biosystems Inc.). Bead size was determined by the nozzle size and further controlled by adjusting vibration frequency and amplitude, as well as flow rate. Optimal parameters for bead formation were achieved by visualization of real-time bead formation as the beads were passed through the nozzle in front of the stroboscope lamp on the encapsulator. That is, beads that were separated for several centimeters and dispersed in a circular flow indicated optimal bead formation. Once these parameters were established they were preset and used consistently throughout all encapsulation 29 experiments. Poorly formed beads occurring at the beginning and end of an encapsulation run were intercepted using the bead bypass collection cup. Optimal beads subsequently dropped into polymerization solution containing calcium-chloride ions that caused the beads to harden. This hardening solution was then drained and the beads were washed with M O P S solution followed by resuspension in H G - D M E M . The beads were then collected in the bead collection flask. RU486 induction of encapsulated versus non-encapsulated leptin-producing gut cells Leptin-producing gut-cells were grown in 10 cm plates until 80-85% confluent. Half of each plate was seeded directly into 24-well plates as in the previous in vitro study and the other half was encapsulated in a 1.5% sodium-alginate coating; encapsulated cells were later transferred to 24-well plates (500 pL of encapsulated cells/well). Forty-eight h later, both the non-encapsulated and encapsulated leptin-producing gut-cells were induced with various doses of RU486 (0 M, 10" 1 4 M, 10" 1 2 M, 10- 1 1 M, 1 0 - 1 0 M , 10" 9 M, 10- 8 M or l O ^ M ) . Media samples were collected 24 h later, centrifuged, and stored at -80 C until further analysis. Human leptin concentrations in the media were determined using the human leptin ELISA kit. RU486 induction of encapsulated leptin-producing gut cells cultured for 5 weeks Leptin-producing gut-cells were grown in a 10 cm plate until 85% confluent then encapsulated in a 1.5% sodium-alginate coating, and resuspended in approximately 100 mL of H G - D M E M . Encapsulated leptin-producing gut cells were maintained in culture with H G - D M E M . At weeks 1, 3 and 5 post-encapsulation, 12 mL of encapsulated cells were removed from culture and 500 pL of these encapsulated cells 30 were pipetted into each well of a 24-well plate. All encapsulated leptin-producing cells were incubated with either no RU486 or with 10"8 M RU486 for 24 h at which time media was collected and centrifuged. The supernatant was transferred to clean 1.5 mL microcentrifuge tubes and stored at -20 C. Human leptin concentrations in the media were determined using the human leptin ELISA kit. Animals Male ob/ob mice were used for the majority of the animal studies and, unless otherwise stated, mice were 5 ± 2 weeks of age. These mice were purchased from Jackson Laboratories (Bar Harbor, MO, USA) and were housed in groups of 5-6 mice, except for the kidney capsule transplant studies where the mice were individually housed. All ob/ob mice had access to standard mouse chow (Formula 5015, 3.73 total kcal per gram of food; Lab Diets, St. Louis, MO, USA) and water ad libitum. For studies involving immunosuppressant administration, ob/ob mice were administered intraperitoneal cyclosporine (Cs; Sandimmune; 15 mg/kg); control animals received saline. Male C57BL6 /J mice, aged 4-5 weeks, were used for the high fat diet studies; these mice were also obtained from the Jackson Laboratories (Bar Harbor, M O , USA) and were housed individually or in small groups of 2-4. They were fed a high fat diet consisting of 58% kcal from cocunut oil fat (Formula D12330, 5.56 total kcal per gram of food; Research Diets, N J , USA) . All animals were housed in a facility that maintained a 12-hour light/dark cycle. Animal work was performed in accordance with and approval from the U B C Animal Care Committee, following the Canadian Council on Animal Care guidelines. 31 RU486 pellets Slow-release RU486 pellets were purchased from Innovative Research of America (Florida, USA) . These pellets were designed to release RU486 over a 14-day period at a constant release rate. Initially, at the time of transplantation of control or leptin-producing gut cells, these RU486 pellets were implanted into the intraperitoneal cavity of mice. In experiments where more than one RU486 pellet was implanted, the consecutive pellets were implanted at the end of the 14-day release of the previous pellet; these consecutive pellets were implanted subcutaneously at the base and dorsum of the neck. The RU486 concentration of pellets used in these studies varied between 0.875 pg/pellet, 8.75 pg/pellet, 35 pg/pellet, or 87.5 pg/pellet (Tables III, IV and V). 32 TABLE III: KIDNEY CAPSULE TRANSPLANT STUDIES IN OB/OB MICE (STUDIES 11.1 AND II.2) Figure Experiment Group Number Age of Concentration Number Transplanted Cyclosporine Number Number of animals of RU486 of RU486 cell type treatment animals (weeks) pellet (pg) pellets (15mg/kg) per group 7 11.1 A 6 > 10 wk 35 1 Leptin B 5 > 10 wk 35 1 control 9 II.2 A 5 10 wk 35 2 Control Yes B 5 10 wk 35 2 Leptin Yes c 5 10 wk 35 2 Leptin No Leptin cell type = leptin-producing gut cells; control cell type = control gut cells TABLE IV: ENCAPSULATION TRANSPLANT STUDIES IN OB/OB MICE (STUDIES 111.1 TO III.7) Figure Experiment Group Number Age of Concentration Number Transplanted Volume of Number Number of animals of RU486 of cell type capsules Animals (weeks) pellet (pg) RU486 transplanted (per pellets (mL) grp) 14 111.1 A 4 4 wks 87.5 1 Control 1 B 4 4 wks 0.875 1 Leptin 1 C 4 4 wks 8.75 1 Leptin 1 D 4 4 wks 87.5 1 Leptin 1 15 III.2 A 6 4 wks 87.5; 8.75 2 Leptin 1.5 B 6 4 wks 87.5 (x4); 8.75 5 Leptin 5 C 6 4 wks 87.5; 8.75 2 Leptin 2.5 D 6 4 wks 87.5 (x4); 8.75 5 Control 2.5 18 III.3 A 1 8 wks 87.5 1 Control 2.5 B 2 8 wks 87.5 1 Leptin 2.5 C 2 8 wks 87.5 1 Leptin 0.25 19 III.4 A 6 5 wks 87.5 2 Control 0.5 B 6 5 wks 87.5 2 Leptin 0.5 C 6 5 wks 8.75 2 Leptin 0.5 22 III.5 A 6 10 wks 87.5 1 Control 0.5 B 6 10 wks 87.5 1 Leptin 0.5 C 6 10 wks 8.75 1 Leptin 0.5 26 III.6 A 6 4 wks 0 0 None 0 B 6 4 wks 87.5 3 Control 2 C 6 4 wks 87.5 3 Leptin 2 Leptin = leptin-producing gut cells; control = control gut cells 33 T A B L E V: E N C A P S U L A T I O N T R A N S P L A N T STUDIES IN HIGH F A T F E D C57BL /6J MICE (STUDIES IV.1 T O IV.5) Figure Experiment Group Number A g e of Concentrat ion Number Transplanted Volume of # cel ls Number Number of animals of RU486 of cell type capsu les per Animals (weeks) pellet (pg) RU486 transplanted capsu le (per pellets (mL) (million) grp) 30 IV.1 A 6 4 wks - 0 0 0 0 B 6 4 wks - 0 0 0 0 C 6 4 wks 87.5 3 Control 5 4.5 D 6 4 wks 87.5 3 Leptin 5 4.5 31 IV.2 A 6 4 wks - 0 0 0 0 B 6 4 wks - 0 0 0 0 C 6 4 wks 87.5 1 Control 5 4.5 D 6 4 wks 87.5 1 Leptin 5 4.5 35 IV.3 A 2 >10 wks — 0 0 0 B 2 >10 wks 87.5 1 Control 5 18 C 2 >10 wks 87.5 1 Leptin 5 18 36 IV.4 A 6 4 wks - 0 0 0 B 6 4 wks 87.5 1 Control 5 18 C 6 4 wks 87.5 1 Leptin 5 18 40 IV.5 A 2 >10 wks 87.5 1 Control 5 18 B 2 >10 wks 87.5 1 Leptin 5 18 Leptin cell type = leptin-producing gut cells; control cell type = control gut cells 34 In vivo studies Body weight and food intake were monitored three times per week for all animal studies, except in the kidney capsule studies where body weight and food intake were monitored daily. Food was initially weighed on day 1, for example. On day 3 the food would be re-measured prior to replenishing the supply. Food consumption was calculated by subtracting the weight of food left in the cage on day 3 from the weight of the food given on day 1. This value would then be divided by the number of days between weighings and would enable the evaluator to calculate the amount of food consumed per cage. Blood glucose was monitored using a One Touch Ultra glucometer (LifeScan Inc., Milpitus, CA) , two times per week and blood samples were collected once per week from the tail vein of conscious, unrestrained mice. For serum collection, blood was obtained from the tail vein of unrestrained, conscious mice into non-heparinized capillary tubes and transferred to 1.5 mL microcentrifuge tubes. Blood was allowed to clot for 30 minutes at room temperature then centrifuged at 7600xg for 9 m; serum was transferred to clean 1.5 mL microcentrifuge tubes and stored a t - 2 0 C. P lasma was obtained by collecting blood from the tail vein into heparinized capillary tubes; blood was transferred to 1.5 mL microcentrifuge tubes containing sodium heparin (50 units). Blood samples were centrifuged at 7600xg for 9 m; plasma was transferred to clean 1.5 mL microcentrifuge tubes and stored at -20 C. Plasma/serum samples were later assayed for human leptin (RIA kits from A L P C O Diagnostics (Windham, USA) ; or L INCO Research Inc. (Missouri, USA) , mouse insulin (ELISA; A L P C O Diagnostics) and triglycerides (kit with modifications based on those by Weber et al. (108); Sigma Aldrich). The A L P C O Diagnostics RIA kit was reported to have only little cross-reactivity with mouse, rat, horse, sheep and chicken leptin but 100% cross-reactivity with human leptin. This A L P C O kit could detect as low as 0.04 ng/mL of 35 leptin. The sensitive human leptin RIA kit from LINCO Research Inc. had 100% cross-reactivity to human leptin and < 1 % cross-reactivity to rat leptin and < 2% cross-reactivity to mouse leptin. The lowest level of leptin that can be detected by this L INCO kit is 0.05 ng/mL of leptin. Transplantation of leptin-producing gut cells under the kidney capsule Control or leptin-producing gut cells were grown to 80-85% confluence prior to transplantation. Cells were washed once with ice-cold Dulbecco's phosphate buffered saline (PBS, Invitrogen Canada Inc.) then incubated with 0.1% trypsin/1 mM E D T A for 7-8 m. P B S (with CaCb) was added to terminate the trypsinization process and the cell suspension was transferred to a 15 mL conical vial. Cells were pelleted by centrifugation at 200xg for 2 m and the supernatant was aspirated and discarded. The cell pellet was resuspended in 1 mL serum-free H G - D M E M and the cell suspension was divided into 500 uL aliquots. This cell suspension was aspirated into a micro-tube, clamped with Liga-clips (Autoclip®, Becton Dickinson) and centrifuged using a specialized micro-tube centrifuge and micromanipulator (a generous gift from Dr. Ray Rajotte, University of Alberta, Edmonton, Canada) to pellet the cells at the end of the micro-tube. Recipient mice were anesthetized with isoflurane and the dorsal side of the mouse was exposed, shaved and wiped down with 70% alcohol. A skin incision was made dorsally in the region of the kidney (1 cm length) and the dermis was separated from the underlying body wall. After making an incision in the body wall, the kidney was located and externalized. The kidney capsule was cut and a glass probe was used to form a small pocket into which control gut or leptin-producing gut cell pellets were expelled from the micromanipulator. An RU486 pellet was implanted into the kidney region and the incisions were closed and sutured. 36 In Experiment 11.1, ob/ob mice (approximately 10 weeks old) were divided into two groups (Table III). Group A (n=6) was transplanted with leptin-producing gut cells plus a 35 pg RU486 pellet and group B (n=5) was transplanted with control gut cells plus a 35 pg RU486 pellet (Table III). Serum collection and body weight, food intake and blood glucose monitoring were as previously described. In Experiment 11.2, ob/ob mice (9-10 weeks old) were divided into three groups (Table III). Group A (n=5) was transplanted with control gut cells plus a 35 pg RU486 pellet and C s (15 mg/kg); group B (n=5) was transplanted with leptin-producing gut cells, a 35 pg RU486 pellet with saline instead of C s . Finally, group C (n=5) was transplanted with leptin-producing gut cells, a 35 pg RU486 pellet and C s (15 mg/kg). C s was administered to ob/ob mice intraperitoneally by injection. Body weight, food intake, blood glucose, and serum leptin were monitored as previously described. Transplantation of encapsulated leptin-producing gut cells or control gut cells Control gut cells or leptin-producing gut cells were grown to 80-85% confluence. Control gut or leptin-producing gut cells were encapsulated as described previously. Unless otherwise stated, for all encapsulation studies, 5 mL, 2.5 mL, 1.5 mL and 500 pL of encapsulated cells was equivalent to approximately 4.5 million cells, 2.25 million cells, 1.4 million cells and 0.45 million cells, respectively. Encapsulated cells were resuspended in P B S without calcium, drained from the encapsulator and drawn into syringes, ensuring equal volumes of capsules per syringe. Intraperitoneal injection of capsules was performed as follows: mice were anesthetized using isoflurane and the ventral side of the mouse was exposed, shaved and wiped down with 70% alcohol. A small skin incision (1 cm) was made between the midline and right side of the animal. Another small incision (0.5 cm) was made in the abdominal muscle wall in the same region. Insertion of an RU486 pellet into the 37 intraperitoneal cavity was completed followed by injection of the capsules through an 18-guage i.v. catheter (BD Insyte™, Becton Dickinson). Depending on the study, various volumes of capsules were injected and different concentrations of RU486 pellets were administered (see Tables V and VI). Lidocaine was administered superficially and the incisions were sutured. In Experiment 111.1, ob/ob mice were divided into 4 groups, each with 4 ob/ob mice per group (Table IV). Mice in group A received 1 mL of encapsulated control gut cells and one 87.5 u,g RU486 pellet. Groups B, C and D all received 1 mL of encapsulated leptin-producing gut cells; group B received one 0.875 uxj RU486 pellet, group C received one 8.75 \ig RU486 pellet, and group D received one 87.5 u.g RU486 pellet. Body weight, food intake and blood glucose measurements were monitored as previously described. In Experiment III.2, ob/ob mice were divided into 4 groups of 6 ob/ob mice each (Table IV). Group A was transplanted with 1.5 mL of encapsulated leptin-producing gut cells. Groups B and C were transplanted with 5 mL and 2.5 mL of encapsulated leptin-producing gut cells, respectively, while group D was transplanted with 2.5 mL of encapsulated control gut cells. All groups of mice received an initial 87.5 u.g RU486 pellet and after the cessation of RU486 release 14 days after its implantation, a lower dose RU486 pellet (8.75 u.g) was implanted into animals. After the cessation of RU486 release from the second pellet, only groups B and D received RU486 pellets (Table IV). Groups B and D each received 87.5 ug RU486 pellets three more times, at 14-day intervals. Body weight, food intake, blood glucose levels, serum leptin, and serum insulin were all monitored as previously described. Experiment III.3 was a pilot study consisting of three different ob/ob mouse groups (Table IV). Group A (n=1) received 2.5 mL of encapsulated control gut cells and 38 a 87.5 pg RU486 pellet. Group B (n=2) received 2.5 mL of encapsulated leptin-producing gut cells and an 87.5 pg RU486 pellet. Group C (n=2) received 250 pL of encapsulated leptin-producing gut cells and an 87.5 pg RU486 pellet. Body weight was monitored as previously described. In Experiment ill.4, ob/ob mice were divided into three different groups (Table IV). Group A received 500 pL of control gut cells and an 87.5 pg RU486 pellet. Group B received 500 pL of leptin-producing gut cells and an 87.5 pg RU486 pellet while group C received 500 pL of leptin-producing gut cells and an 8.75 pg RU486 pellet. Each group of mice received a second 87.5 pg RU486 pellet 14-days after the initial transplantation. Experiment III.5 followed the same protocol outlined for Experiment 111.4 except using older ob/ob mice (age 10 weeks at the start of the study) and implanting only one RU486 pellet (Table IV). All groups received 500 pL of encapsulated cells, but only received one RU486 pellet (87.5 pg) each. Body weight, food intake, blood glucose, plasma leptin, plasma insulin, and plasma triglyceride levels were measured as previously described. Experiment III.6 consisted of three groups of 6 ob/ob mice each (Table IV). Group A did not receive transplanted cells or RU486 pellets. Group B received 2 mL of encapsulated control gut cells with an initial 87.5 pg RU486 pellet. Group C received 2 mL of encapsulated leptin-producing gut cells with an initial 87.5 pg RU486 pellet. Groups B and C also received 2 other 87.5 pg RU486 pellets at 14-day intervals from implantation of the first RU486 pellet. Body weight, food intake, blood glucose concentrations, serum leptin, and serum triglyceride levels were measured, as previously described. 39 In experiment IV.1, C57BL/6J mice were divided into 4 different groups (n=6) such that the starting body weights were the same at the start of the study (Table V). Body weight measurements were made, as previously described, for 26 days before mice were either placed on a high-fat diet (HFD) or maintained on a regular chow diet (LFD). Mice from groups A and B did not receive any cell transplants or RU486 pellets and were fed a HFD and L F D , respectively. Mice in groups C and D were also fed a H F D and group C received 5 mL of encapsulated control gut cells with an 87.5 u,g RU486 pellet, while mice from group D received 5 mL of leptin-producing gut cells with an 87.5 ng RU486 pellet. Five mL of encapsulated leptin-producing or control gut cells was equated to approximately 4.5 million cells in this experiment. Groups C and D received two additional 87.5 u.g RU486 pellets, at 14-day intervals from the initial RU486 pellet implantation. Experiment IV.2 also consisted of 4 groups of 6 C57BL/6J mice each (Table V). Groups A and B were as for experiment IV. 1, where neither group received any cell transplants or RU486 pellets; group A was fed a HFD while group B was maintained on a LFD. Groups C and D were also fed a HFD for the duration of the study. When the body weights of groups A, C and D were significantly higher than that of group B (P<0.05), transplantation of leptin-producing gut or control gut cells was performed. Mice from group C received 5 mL of control gut cells with an 87.5 ng RU486 pellet; mice from group D received 5 mL of leptin-producing gut cells with an 87.5 |j,g RU486 pellet. Again, in this experiment, transplanting 5 mL of encapsulated cells was equivalent to transplanting approximately 4.5 million leptin-producing or control gut cells per animal. Body weight, food intake, blood glucose concentrations, serum leptin, serum insulin, and serum triglyceride levels were measured, as previously described. 40 Experiment IV.3 was a pilot study where 6 C57BL/6J mice were divided into 3 groups of 2 each (Table V). Group A was maintained on a H F D but did not receive any cell transplant or RU486 pellet. Group B received 5 mL of encapsulated control gut cells with an 87.5 pg RU486 pellet. Mice from group C received 5 mL of encapsulated leptin-producing gut cells with an 87.5 pg RU486 pellet. In this experiment, 5 mL of encapsulated cells equated to approximately 18 million leptin-producing or control gut cells. Ten days after transplantation of the leptin-producing or control gut cells, all animal groups received an intraperitoneal injection of exogenous RU486 (250 pg/kg body weight). Body weight was monitored three times per week as previously described. In experiment IV.4, C57BL/6J mice were divided into three groups of 6 each (Table V). All mice were maintained on a H F D . Group A did not receive any cell transplant or RU486 pellet and represented the HFD-control group. Group B received 5 mL of encapsulated control gut cells (-18 million cells) with an 87.5 pg RU486 pellet. Group C received 5 mL of encapsulated leptin-producing gut cells (-18 million cells) with an 87.5 pg RU486 pellet. Body weight, food intake, blood glucose, serum leptin, serum insulin and serum triglyceride levels were monitored as previously described. Experiment IV.5 was the final pilot study performed using 2 groups of 2 C57BL /6J mice each (Table V). All mice were maintained on a HFD. Leptin-producing and control gut cells were encapsulated 1 week prior to transplantation. Twenty-four h before transplantation, all encapsulated cells were incubated with 10"8 M RU486 to induce the leptin-producing gut cells to produce leptin prior to transplantation. Group A received 5 mL of encapsulated control gut cells (approximatelyl 8 million cells) and an 87.5 pg RU486 pellet. Group B received 5 mL of encapsulated leptin-producing gut cells 41 (approximately18 million cells) and an 87.5 pg RU486 pellet. Body weight was measured 3 times per week, as previously described. DATA ANALYSIS Data are presented as mean±standard error of the mean (SEM), mean + S E M or as the mean. Statistical significance was assessed using a student's t-test or analysis of variance (ANOVA). Whenever A N O V A was used, it was followed by either Dunnett's Multiple Comparison or Newman-Keuls post hoc tests to further evaluate the significance between groups. Statistical significance was set at the 5% confidence interval where *P<0.05, **P<0.01, and ***P<0.001. 42 RESULTS STUDY I: •IN VITRO CHARACTERIZATION OF LEPTIN-PRODUCING GUT CELLS 1.1: GIP mRNA expression Leptin-producing gut cells were genetically engineered by previous members of the Kieffer lab, at the University of Alberta, by transforming GTC-1 cells into two different cell lines: GTC-1 pSwitchGIPro (control gut cells) and G T C -IpSwitchGIProhLeptin (leptin-producing gut cells) (Figures 1 and 2). GTC-1 cells are a gut endocrine cell line which normally express and release the incretin hormone GIP (98, 107); the engineered GTC-1 cells were designed to produce and release leptin in response to Mifepristone (RU486). To determine if GIP production was altered by the engineering of these cells, GIP m R N A expression levels were compared in GTC-1 cells, control gut cells (GTC-1 pSwitch cells) and leptin-producing gut cells (GTC-IpSwitchGIProhLeptin cells) using Northern blot analysis (Figure 3 A). There was no significant difference in relative GIP m R N A expression in GTC-1 cells as compared to control gut cells or leptin-producing gut cells (121% versus 127% and 125%, respectively, as normalized to B-actin). Data from the slot blot analysis confirmed that there was no difference in relative GIP mRNA expression when comparing the 3 cell lines (Figure 3 B). 43 A. B. GIP P-Actin N o R U 4 8 6 1 0 - U M R U 4 8 6 40 pg R N A 20 pg R N A GTC-1 cells 40 ug R N A 20 ug R N A Control gut cells 40 pg R N A 20 ug R N A Leptin-producing gut cells F igure 3. GIP m R N A expression in GTC-1 cells compared to control gut cells (GTC-1 pSwitchGIPro cells) and leptin-producing gut cells (GTC-1 pSwitchGIProhLeptin cells). (A) Upper blot shows GIP m R N A expression while bottom blot shows p-actin. Densitometric values for GIP m R N A levels are expressed as % of corresponding p-actin values. (B) Slot blots show GIP m R N A expression between the three different cell lines, in the absence or presence of RU486. 44 1.2: Leptin mRNA expression Preliminary experiments conducted at the University of Alberta examined whether the engineered cells containing the leptin gene did, in fact, express leptin m R N A and further, whether leptin m R N A levels changed in a dose- and time-dependent manner in response to RU486. Northern blot analysis demonstrated that leptin m R N A expression increased in leptin-producing gut cells in an RU486 dose- and time-dependent manner (Figure 4). Treatment of the leptin-producing gut cells with 10" 1 2 M RU486 for 24 h resulted in a 2.3±1.5-fold increase in human leptin m R N A expression as compared to leptin-producing gut cells that were not stimulated with RU486. Treatment of leptin-producing gut cells for 24 h with 10" 1 0 M, 10" 8 M, or 10"6 M RU486 resulted in a 18.3±1.2, 64.7±5.7 and 61.3±9.1 fold increase (respectively) over leptin-producing gut cells receiving 10" 1 2 M RU486 stimulation. No detection of leptin m R N A was observed in the absence of RU486. When leptin-producing gut cells were induced with 10"8 M RU486 for 0 h, 4 h, 8 h and 24 h, the amount of human leptin mRNA expressed showed a 3.0+1.0, 43.3+7.7, 56.3±6.7 and 77.7±6.7 fold increase over leptin-producing gut cells without RU486 stimulation, respectively. Minimal levels of leptin m R N A were detected in the absence o f R U 4 8 6 . 1.3: Leptin Protein Expression Western blot analysis of leptin protein expression was conducted at the University of Alberta. The amount of leptin protein expression observed after induction with 1 0 " 1 2 M , 1 0 - 1 0 M , 10 - 8 M, o r l O ^ M R U 4 8 6 w a s 1.0±0,10.7±1.5, 42.7±0.7 and 52.7±5.2 fold increase over 10" 1 2 M RU486, respectively (Figure 5). Leptin protein expression was also assessed at 0 h, 4 h, 8 h and 24 h after induction with 10"8 M 45 RU486 and was found to be 1.0±0, 7.7±4.2, 34.3±1.9 and 62.7±3.2 fold increase over control, respectively. Leptin was thus expressed in an RU486 dose- and time-dependent manner and minimal leptin protein was detectable in the absence of RU486. 1.4: Leptin Protein Release Leptin protein release from the leptin-producing gut cells was evaluated at the University of Alberta by ELISA (Figure 6). The amount of leptin released increased from 1.9±0.5 ng/mL at time zero to 667.8±130.3 ng/mL after incubating for 24 hours with 10"8 M RU486. Cel ls were also induced with 10" 1 2 M, 10" 1 0 M, 10 - 8 M or 10 - 6 M RU486 and the amount of leptin secreted at these various RU486 concentrations was: 2.0+0.4 ng/mL, 26.7±6.5 ng/mL, 280.0±106.6 ng/mL and 250.3±71.2 ng/mL respectively. These results indicate that leptin is released from these genetically engineered gut cells in a time- and dose-dependent manner in response to RU486. 46 A. Time (h) 1 ~ ol. • I U U £ 10" 1 2 1 0 1 0 1 0 ' 8 1 0 6 RU486 Dose (M) Figure 4. Leptin m R N A expression in leptin-producing gut cells. Leptin-producing gut cells incubated with RU486 for various time points (A) and incubated with various concentrations of RU486 (B). Densitometric leptin m R N A levels expressed as fold increase over 0 h or 10 " 1 2 M RU486. n=3; *P<0.05 (comparing all groups to 0 h or 10 " 1 2 M RU486). A. 47 c _ 7 0 § ? 60 * •_ 40-LU c '5 * J 2 C L C 0 E 30 § 2 0 -« 0 4 8 Time (h) 24 B. c 6O-1 g § 501 g->40-L U .S .E X 30-1 5 g S 20-— w a © 10-Q- Q © —• 10 12 1 Q . 1 0 1 Q - 8 RU486 Dose (M) 10 -6 Figure 5. Leptin protein expression in leptin-producing gut cells. Leptin-producing gut cells were incubated with RU486 for various time points (A) or incubated with various doses of RU486 (B). Protein lysates from leptin-producing gut cells were immunoblotted with anti-hLeptin antibodies. Representative immunoblots depicting leptin protein expression are shown above each bar graph. Bar graphs show the densitometric analysis of leptin protein expression. n=3; *P<0.05, **P<0.01 (comparing 0 h to 4, 8 and 24 h; comparing 10" 1 2to 10"1 0,10"8 and 10"6 M RU486). 48 B. 2 5 0 0 -_ 2 0 0 0 -_ i E o) 1500 -c » 1000 -Q. o 5 0 0 -o -** ** ** X *** X ** X 24.0 0.0 4.0 8.0 24.0 32.0 48 .0 56.0 Time (h) 700- i 600-1 ^ 5 0 0 g>400-| £ 300H § " 2 0 0 - 1 100-1 0 I I 1 0 - 1 6 1 0 - 1 4 1 0 - 1 2 1 0 - 1 1 1 0 - 1 0 1 0 " 9 10" 8 10" 6 D o s e o f R U 4 8 6 (M) Figure 6. Leptin protein release from leptin-producing gut cells. Leptin-producing gut cells incubated with RU486 after various time points (A) and incubated with various doses of RU486 (B), measured by ELISA. n=4; *P<0.05, **P<0.01, ***P<0.001 (comparing all time points to 0 h and all dose levels to 1 0 1 6 M RU486). 49 STUDY II: TREATMENT OF OBESITY AND DIABETES IN OB/OB MICE BY TRANSPLANTATION OF LEPTIN-PRODUCING GUT CELLS UNDER THE KIDNEY CAPSULE 11.1: Effects of the transplantation of leptin-producing gut cells under the kidney capsule of ob/ob mice Body weight, food intake and blood glucose Pre-transplant body weights were not significantly different between groups A (control) and B (leptin-treated) (Figure 7). By day 13 post-transplantation, group B had reached its lowest body weight (53.4±2.6 g) and weighed significantly less than group A (59.1+0.9 g (P<0.05)). Daily food intake data also indicated a significant decrease in food intake from day 1 post-transplantation to day 15 post-transplantation in the leptin-treated group (group B, Figure 7). Average food intake prior to the transplantation was 8.9±0.2 g and 8.3±0.9 g for groups A and B respectively. Maximum decreases in food intake were noted at day 8 post-transplantation where group B consumed 3.5+0.3 g while group A consumed 7.3±0.4 g of food (PO.001) . By day 15 post-transplantation, food intake of the leptin-treated group (group B) had gradually increased to 5.9+0.6 g but remained significantly lower than the control group A, 7.1 ±0.3 g (P<0.05). Body weight and food intake values started to increase approximately 13 days post-transplantation even though RU486 was still being released. Prior to transplantation, blood glucose levels for group B were 23.0+2.3 mM and 27.0±1.3 mM for group A (Figure 7). By seven days after transplantation, blood glucose levels had dropped to 8.0±0.3 mM in the leptin-treated group B, but were 24.5±4.3 mM 50 in the control group A, (PO.01 ) . Average blood glucose levels remained significantly different between groups A and B up to 38 days post-transplantation, where the leptin-treated group B remained normoglycemic while group A remained hyperglycemic. These decreased blood glucose levels were observed in the leptin-treated group B despite the fact that body weight had returned to pre-transplant levels by day 17 post-transplantation. 51 704 — 65 a I 60 > 1 00 55 50 TT T T X ! • Leptin-producing gut-cells Control gut cells B. — i — 10 Transplant 20 30 Duration (days) - 1 — 40 — i — 50 — i — 60 12.5 _*10.0 at 5 7.5-•a 5.0 o o 2.5-0.0 RU466 pellet 1 RU486 pellet 2 ^ c. —I— 10 Transplant 20 — i — 30 40 50 — i — 60 Duration (days) S § I o o o •a o o m 35 30 25 20 15 10 5 RU486 pellet 1 RU486 pellet 2 ^ — i — 10 — i — 20 — i — 30 - i — 40 50 60 Duration (days) Figure 7. Transplantation of control gut or leptin-producing gut cells under the kidney capsule of ob/ob mice. Body weight (A), food intake (B) were monitored 3 days per week and blood glucose concentrations (C) were monitored 2 days per week, in ob/ob mice. Ob/ob mice received leptin-producing gut cells (blue • , n=4) or control gut cells (orange a, n=4) to their kidney capsule at the same time that they received a 35 pg RU486 pellet. A second RU486 pellet was given 14 days later. All values plotted as mean±SEM. *P<0.05, ** P<0.01, ***P<0.001. 52 Circulating human leptin levels In order to confirm whether the leptin-producing gut cells, transplanted under the kidney capsule of ob/ob mice, were secreting human leptin, serum human leptin was measured using a commercially available RIA kit (LINCO). Preliminary measurements of serum human leptin levels in these ob/ob mice revealed inconsistent and inconclusive leptin levels in the control and the leptin-treated groups (data not shown). Human leptin concentrations measured in the leptin-treated group were not significantly different from those measured in the control group. It was hypothesized that the inconsistent data might be due to interference with leptin-binding proteins present at high concentrations in the ob/ob mouse serum. Hence, a comparison study was conducted to measure serum and plasma leptin levels in the same samples using two different commercially available RIA kits (Figure 8). Data obtained using a L INCO human leptin RIA kit were compared to data obtained using an A L P C O human leptin RIA kit. All ob/ob serum and plasma samples were from animals that had not received any leptin-producing gut or control gut cell transplants, thus we were anticipating immunoreactive leptin levels of close to zero. The A L P C O kit did not detect any human leptin in the ob/ob plasma samples but detected up to 2-2.5 ng/mL of human leptin in the ob/ob serum samples. The LINCO kit detected up to 0.5 ng/mL human leptin in both ob/ob plasma and serum samples. As well, ob/ob plasma and serum samples were spiked with recombinant leptin prior to being assayed and both the A L P C O and L INCO RIA kits detected the expected concentration of spiked leptin, taking into account the background values (data not shown). Interference with leptin antibody binding was detected using both RIA kits but less interference with ob/ob serum samples was noted using the LINCO kit versus the A L P C O kit; little to no interference with ob/ob plasma samples was noted using the A L P C O kit. Therefore, for subsequent 53 assays, we decided to use the LINCO human leptin RIA kit to measure ob/ob serum leptin levels and the A L P C O human leptin RIA kit to measure ob/ob plasma samples. 54 B. Figure 8. Human leptin radioimmunoassays: comparison of results obtained from an A L P C O human leptin RIA (A) and a L I N C O human leptin RIA (B). P lasma and serum samples from ob/ob mice, not treated with leptin cell therapy, were assayed on each RIA to determine differences in the level of non-specif ic binding of the hLeptin antibody to determine background levels circulating in ob/ob p lasma or serum. Values plotted as mean+SEM. ***P<0.001 (comparing ob/ob p lasma or serum with the leptin antibody added to ob/ob p lasma or serum with no leptin antibody added). 55 11.2: Effects of the transplantation of leptin-producing gut cells to the kidney capsule of ob/ob mice receiving immunosuppressant therapy Body weight, food intake and blood glucose We hypothesized that the short-term efficacy of leptin treatment on body weight and food intake seen in study 11.1 was due to immunodestruction of the transplanted leptin-producing cells. In order to determine whether this was the case, we designed another experiment wherein ob/ob mice received transplant of control or leptin-producing cells under their kidney capsule, with or without immunosuppressant therapy (cyclosporine, Cs) (Figure 9). Pre-transplant body weights were not significantly different (60.2±1.6 g, 61.3±0.6 g and 60.9±1.7 g for groups A, B, and C respectively). All groups initially lost weight due to surgical stress; however, by day 15 post-transplantation, the group that received leptin-producing gut cells with RU486 and C s (group C) had significantly lower body weight than controls (53.7±2.0 g (group C) versus 57.6±2.1 g and 60.9±0.4 g for groups A and B respectively; P<0.05). Group B, which received the leptin-producing gut cells and RU486 without C s , started regaining body weight within 7 days post-transplantation such that by day 15 post-transplantation their body weight was almost the same as their pre-transplant body weight (60.8±0.4 g). Group A received control gut cells, RU486 and C s and, although these ob/ob mice were anticipated to gain weight, their body weight decreased at a rate of -0.4±0.1 g/day from day 17 post-transplantation to the end of the study. During this same time, body weight gain was 0.15±0.03 g/day and 0.45±0.06 g/day in groups B and C, respectively. On the day of transplantation, food intake for groups A , B and C was 7.5±0.8 g, 7.5±0.8 g and 6.2±0.7 g respectively, which were not significantly different from one another (Figure 9). The food intake decreased to 6.0+0.5 g, 5.6+0.3 g and 4.7±0.6 g for 56 groups A, B and C by day 4 post-transplantation. Eleven days post-transplantation, food intake for groups A, B and C was 6.9±0.6 g, 6.1 ±0.9 g and 4.4±0.6 g (P<0.05, comparing group C to groups A and B). No significant differences in food intake were observed between the three groups after day 11 post-transplantation. Blood glucose levels were not significantly different from one another prior to transplantation: 23.7±4.6 mM (group A), 26.4±5.2 mM (group B) and 19.4±4.1 mM (group C) (Figure 9). On day eleven post-transplantation blood glucose levels were 21.5±4.6 mM (group A), 13.6±3.8 mM (group B) and 9.1+0.8 mM (group C) . By day 15 post-transplantation, blood glucose levels were 22.8±5.8 mM, 13.5+2.3 mM and 10.9±0.8 mM for groups A, B and C respectively. Group C responded with a maximum reduction of 65% in blood glucose concentration on day 18 post-transplantation compared to pre-transplant levels; this decrease in glycemia was significantly different from groups A and B (P<0.05) and this persisted throughout the rest of the study, up to almost 40 days after the transplantation. Again, the decreased blood glucose levels remained despite the fact that these mice rapidly regained their body weight such that they were as heavy as their pre-transplant diabetic weight by day 29 post-transplantation. 57 ~ 7 0 -— | SO-T3 o Transplant (day 32) A Control + C s • B: Leptin - C s - C: Leptin + C s I RU486 pellet 1 | RU486 pellet 2 50H —i— 10 —r~ 20 — i — 30 i 40 — i — 50 i 60 Duration (days) ~70 B. 10-, a 8-a I 6H — i — 10 Transplant (day 32) RU486 pellet 1 IRU486 pellet 2 , ~20~ — i — 30 — i — 40 — i — 50 Duration (days) — i — 60 70 Transplant (day 32) | RU486 pellet 11RU486 pellet 2 OH 1 1 1 1 1 1-0 10 20 30 40 50 60 Duration (days) Figure 9. Kidney capsule transplantation of control gut or leptin-producing gut cells into ob/ob mice, with or without Cyclosporine (Cs). Group A (Control + C s , orange • ; n=5) received control gut cells, 35 pg RU486 pellets and C s . Group B (Leptin - C s , blue • ; n=5) received leptin-producing gut cells, 35 pg RU486 pellets and saline. Group C (Leptin + Cs , green • ; n=5) received leptin-producing gut cells, 35 pg RU486 pellets and C s . (A) Body weight, (B) food intake and (C) blood glucose concentrations plotted as mean±SEM. *P<0.05, ** P<0.01, a=P<0.05 (group B versus C), b=P<0.01 (group B versus C), c=P<0.01 (group A versus B), d=P<0.05 (group A versus C). 58 Circulating human leptin levels We first confirmed that control animals were not producing human leptin levels by assaying serum samples for human leptin (Figure 10). Human leptin levels, 2 weeks after transplantation, were 0.1±0.1 ng/mL in the control group A (control gut cells with Cs) , 0.49±0.14 ng/mL in group B (leptin-producing gut cells without Cs) , and 0.71±0.17 ng/mL in group C (leptin-producing gut cells with C s ; P<0.05). Three weeks after transplantation, human leptin levels in the serum were 0.2±0.1 ng/mL (group A), 0.08±0.08 ng/mL (group B) and 0.2±0.1 ng/mL (group C), which were not significantly different from one another. These data indicate that the group C mice receiving control gut cells, RU486 pellets and C s did not secrete human leptin yet longterm body weight loss was observed from day 15 post-transplantation to the end of the study (Figure 9). Ob/ob mice that received leptin-producing gut cells with C s and ob/ob mice that received control gut cells with Csboth experienced body weight loss at different times during the study but visually were very different from one another. Skin integrity and grooming were impaired dramatically in the non-leptin treated group while leptin-treated animals looked healthier and better groomed (Figure 11). 59 0.9-i 0.8-=r0-7-E 0.6-g0.5-c 0.4-Q. 0.3-- 1 0.2-0.1-0.0--6^  0.9-j 0.8--0.7-E, 0.6-g0.5-^0.4-F igu re 10. Serum human leptin levels from ob/ob mice that received leptin cell therapy via kidney capsule transplant, with or without Cyclospsor ine (Cs). Human leptin concentrations measured (A) 2 weeks post-transplantation and (B) 3 weeks post-transplantation; values plotted as mean+SEM. n=5, *P<0.05 (comparing all groups to control cells + C s group). Figure 11. Pho tog raph of an ob/ob m o u s e that r ece ived leptin-p roduc ing gut ce l l s to the k idney c a p s u l e with C y c l o s p o r i n e (Cs) t reatment. 61 STUDY III: TREATMENT OF OBESITY AND DIABETES IN OB/OB MICE BY TRANSPLANTATION OF ENCAPSULATED OF LEPTIN-PRODUCING GUT CELLS 111.1: In vitro leptin secretion from leptin-producing gut cells encapsulated in a 1.5% sodium-alginate polymer coating Encapsulation, as outlined in the Materials and Methods section, allows researchers to envelop cells in a sodium-alginate polymer coating through which cellular products can be secreted, but which is impermeable to immune cells (Figure 12). Thus cells are provided with some protection from immune attack without the use of immunosuppressive drugs. An in vitro experiment was conducted to compare the amount of leptin being secreted by non-encapsulated leptin-producing gut cells and the amount of leptin being secreted from encapsulated leptin-producing gut cells (Figure 13). Non-encapsulated cells were treated with 10 - 1 2 M, IO - 1 0 M, IO - 9 M, 10" 8 M or IO - 6 M RU486 and the amount of leptin secreted at these various RU486 concentrations was: 1.0+0.1 ng/mL, 3.3±0.0ng/mL, 115.6±26.7 ng/mL, 88.1±11.5 ng/mL and 67.3±28.3 ng/mL respectively (Figure 13A). A comparable number of encapsulated leptin-producing gut cells were also treated with the same concentrations of RU486; they released 1.5±0.4 ng/mL, <0.2 ng/mL, 59.0±18.4 ng/mL, 40.8±10.7 ng/mL and 50.5±8.1 ng/mL, respectively (Figure 13 B). The amount of leptin secreted from the encapsulated cells was over 50% less than that secreted by the non-encapsulated cells after treatment withIO"9 M, 10" 8 M and 10"6 M RU486. Although there was a decreased amount of leptin secretion measured in the non-encapsulated cells versus the 62 encapsulated cells, a similar RU486 dose-dependent leptin release profile was observed. Leptin Figure 12. Encapsulation of cells in a sodium-alginate polymer coating. Figure adapted from NovaMatrix Ultrapure Polymer systems (www.novamatrix.biz). 63 A. 15(H E 100H U) c o. 50 X 0 10'12 10"10 10"9 10"8 10"6 RU486 (mM) B. c Q. o 80 70-60-50 40 30 20-10-0 I X io-12 io-10 io*9 <\6* id*6 RU486 (mM) Figure 13. Leptin protein secretion from (A) non-encapsulated or (B) encapsulated leptin-producing gut cells. Non-encapsulated or encapsulated leptin-producing gut cells incubated with various doses of RU486; leptin secretion measured by ELISA. n=4; *P<0.05, **P<0.01, ***P<0.001 (comparing all groups to 0 M RU486 controls). 64 III.2: Transplantation of encapsulated leptin-producing gut cells into ob/ob mice also receiving RU486 pellets of various doses Body weight, food intake and blood glucose At the University of Alberta, leptin-producing gut cells were cultured and then encapsulated in a 1.5% sodium-alginate polymer prior to transplantation into the intraperitoneal cavity of ob/ob mice. Pre-transplant body weights were not significantly different from one another (groups A, B, C and D weighed 41.1±1.4 g, 40.4±3.2 g, 40.4±1.0 g and 40.9±2.2 g, respectively (Figure 14)). On day 13 post-transplantation, control group A had a body weight of 40.5±1.9 g while the three leptin-treated groups (B, C , and D) weighed 33.7±1.8 g, 34.5±0.9 g and 34.6±1.9 g, respectively, which was up to 17% of their pre-transplant body weight. After this time point, these three groups started to regain weight at a rate of 0.48±0.01 g/day (group B), 0.53±0.01 g/day (group C) and 0.48+0.01 g/day (group D) until they had exceeded their pre-transplant body weight by the end of 40 days post-transplantation (rate of body weight gain of control was 0.24±0.01 g/day). All three leptin-treated groups (B, C and D) gained weight at a significantly higher rate than the control group A (P<0.001). Food intake data showed that the control group A had a reduction in food intake from 7.4±0.8 g of chow/day prior to transplantation to 4.6±0.4 g of chow/day by day 6 post-transplantation (Figure 14). Groups B, C and D had larger reductions in food intake over the same time period: group B consumed 6.7+1.1 g of chow/day prior to transplantation and 1.8+0.6 g of chow/day 6 days after surgery. Groups C and D consumed 7.7±0.5 g of chow/day and 6.5±1.2 g of chow/day before transplantation, while they consumed 2.3±0.6 g of chow/day and 2.4±0.4 g of chow/day (respectively), 6 65 days after surgery. Leptin-treated groups (B, C, and D) all consumed significantly less than the control group at this time point (PO.05 ) . One day prior to transplantation, all animals were hyperglycemic (Figure 14). Blood glucose concentrations were 22.5±1.8 mM (group A), 18.0+3.6 mM (group B), 21.0±2.9 mM (group C), and 22.6±1.8 mM (group D). On day 13 post-transplantation, control group A remained hyperglycemic at 26.6±1.5 mM while the three leptin-treated groups were normoglycemic: 7.4+0.4 mM (group B), 7.2±0.4 mM (group C) and 7.5+0.4 mM (group D). Blood glucose concentrations in these leptin-treated 06/06 mice remained reduced by approximately 70% for up to 50 days post-transplantation. 66 5 0 4 5 f " 1 i o CQ 25H 2 0 1 5 Transplant (Day 33) RU486 release , Group A: No leptin-producing gut cells • 87.5 M 9 RU486 —*— Group B: Leptin-producing gut cells + 0.875 ng RLM86 —*— Group C: Leptin-producing gut cells • 8.75 ug RU486 > Group D: Leptin-producing gut cells + 87.5 M 9 RU486 1 0 2 0 3 0 B 4 0 Durat ion (days) 5 0 6 0 7 0 8 0 15H 1<H •o Transplant (Day 33) RU486 release . 1 0 2 0 3 0 40 Durat ion (days) 5 0 6 0 7 0 8 0 4 0 E 3 0 2 0 1 m 1 0 Transplant (Day 33) RU486 release , 1 0 2 0 3 0 4 0 Durat ion (days) 5 0 6 0 7 0 8 0 Figure 14. The effect of leptin cell therapy on (A) body weight, (B) food intake, and (C) blood glucose in ob/ob mice. Groups B (green • , n=4), C (purple T , n=4) and D (blue • , n=4) were transplanted with 1 mL of encapsulated leptin-producing gut cells and received 0.875 pg, 8.75 pg and 87.5 pg RU486 pellets, respectively. Control group A (orange •. n=4) received 1 mL control gut cells and 87.5 pg RU486 pellet. All leptin treated groups experienced transient weight loss (up to 70% of pre-transplant body weight) but maintained longterm normoglycemia. Values plotted as mean + S E M . *P<0.05, **P<0.01, ***P<0.001 (comparing groups B, C and D to A); #=P<0.05 (comparing groups C and D to A). 67 III.3: Transplantation of various volumes of encapsulated leptin-producing gut cells into the intraperitoneal cavity of ob/ob mice also receiving a constant dose ofRU486 This previous study showed that transplant of leptin-producing gut cells in ob/ob mice could induce transient reductions in food intake and body weight yet longer term reductions in blood glucose. It was evident from this that these animals did not lose weight in an RU486 dose-dependent manner, perhaps because the volume of cells transplanted was enough to elicit a maximum body weight loss even with administration of the lowest dose of RU486 (0.875 \ig). The goal of this subsequent experiment was to administer the same concentration of RU486 to all leptin-treated animal groups but vary the volume of encapsulated cells being transplanted. Body weight, food intake and blood glucose Pre-transplant body weights were not significantly different between groups A, B, C and D (53.7±1.1 g, 53.6±0.7 g, 52.4+1.4 g and 53.0+0.8 g, respectively; Figure 15). Fourteen days post-transplantation, when the first 87.5 pig RU486 pellet expired, all leptin-treated groups had responded with similar maximum body weight reductions. Body weights at this time for each group were 47.9±1.7 g (group A), 47.1 ±0.5 g (group B), 48.8±1.3 g (group C) and 55.8±1.2 g (group D, control). Body weight values amongst these 4 groups were statistically different (P<0.001, comparing group D with groups A, B and C). Group B, receiving the highest volume of encapsulated leptin-producing gut cells (5 mL), responded with the greatest weight loss (6.5±0.2 g). Groups A and C received smaller volumes of leptin-producing gut cells (1.5 mL and 2.5 mL, respectively) and lost 5.8±0.6 g (group A) and 3.6±0.1 g (group C) . Although no significant differences in body weight reductions were observed between the three 68 leptin-treated groups, there appeared to be a trend towards a dose-dependent weight loss. After the second, 8.75 ug RU486 pellet was implanted, the body weights of all ob/ob mouse groups increased at similar rates (Figure 15). Body weights for groups A , B and C at the end of RU486 release from the 2 n d pellet (day 28 post-transplantation) were 50.7±1.5 g, 49.1 ±1.6 g, and 53.0±1.3 g, respectively. The body weight for control group D was 61.9±1.4 g. Groups B and D received 3 more 14-day release 87.5 \ig RU486 pellets (on days 28, 42 and 56 post-transplantation) (Figure 15). Group B was the only leptin-treated group to receive this pellet as this group had responded optimally out of the three leptin-treated groups and we wanted to determine whether this group would continue to have either further body weight loss or maintenance of reduced body weights. Body weight from day 16 post-transplantation to the end of the study, however, continued to increase such that the rate of body weight gain for groups A, B, C and D were 0.29+0.02 g/day, 0.24±0.02 g/day, 0.30±0.02 g/day and 0.19±0.02 g/day, respectively. On the day of transplantation, pre-transplant food consumption for groups A, B, C and D was 7.5±0.4 g, 7.7±1.2 g, 7.7±1.2 g and 7.7±1.2 g, respectively (Figure 15). At the end of the first 87.5 u.g RU486 pellet release, food intake values were 2.4±0.1 g (group A), 1.8±0.2 g (group B), 3.0±0.1 g (group C), and 4.5±0.4 g (group D). After implantation of the second 8.75 \ig RU486 pellet, food intake increased at a rate of 0.6±0.1 g of food per day (group A), 0.8±0.1 g of food per day (group B), 0.48±0.0T g of food per day (group C), and 0.2±0.0 g of food per day (group D). After the third, 87.5 u.g RU486 pellet was implanted into groups B and D, the rate of food intake for control group D was 0.25±0.3 g of food per day while the rate of food intake for group B was 69 0.2±0.1 g of food per day. These data indicate that after the implantation of the second RU486 pellet the cells were likely no longer releaseing sufficient leptin to reduce food intake in the leptin-treated groups (A, B and C). Pre-transplant blood glucose concentrations measured 5 days prior to transplantation, were 31.4±1.4 mM (group A), 26.8±2.4 mM (group B), 28.1 ±2.4 mM (group C), and 25.6±2.0 mM (group D). On day 9 post-transplantation, blood glucose concentrations decreased in the leptin-treated groups: 9.4±1.2 mM (group A), 8.0±0.7 mM (group B), and 9.8±0.5 mM (group C); blood glucose values for the control group at this time were 24.4+2.5 mM. Blood glucose levels remained significantly lower in the leptin-treated groups up to day 28 post-transplantation; thereafter, no further significant differences were observed comparing the 3 leptin-treated groups (A, B and C) to the control group (D). 70 Transplant Day RU486 pellet 1 ( 8 7 . 5 M g ) - A: leptin cells ( 1 5 mL capsules) - B: leptin cells ( 5 . 0 mL capsules) - C leptin cells ( 2 5 mL capsules) D: control cells ( 2 . 5 mL capsules) R U 4 8 6 pellet ( 8 . 7 5 ng) R U 4 8 6 pellets 3 & 4 ( 8 7 . 5 u g ) Groups B and D only B . 4 0 SO D u r a t i o n ( d a y s ) 110 Transplant Day R U 4 8 6 pellet 1 ( 8 7 5 ng) R U 4 8 6 pellet 2 R U 4 8 6 pellets 3 & 4 ( 8 7 5 ug) ( 8 7 5 ug) Groups B and D only f 1 ' 10 20 30 4 0 50 60 70 8 0 D u r a t i o n ( d a y s ) Transplant Day R U 4 8 6 pellet 1 ( 8 7 . 5 ( ig ) 90 100 110 ° 0 10 20 30 4 0 50 60 70 80 90 100 110 D u r a t i o n ( d a y s ) Figure 15. The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose in ob/ob mice transplanted with various volumes of cells. Groups A (green • , n=6), B (blue T , n=6) and C (purple • , n=6) received 1.5 mL, 5.0 mL and 2.5 mL encapsulated leptin-producing gut cells to their i.p. cavity, respectively. Group D (orange • , n=6) received encapsulated control gut cells. Each group of mice received RU486 pellets at different times throughout the study. Values plotted as mean ± S E M . *P<0.05, **P<0.01, ***P<0.001 (groups A, B and C to D); °°P<0.01 (groups A and B versus D); $P<0.05 (group B versus D); /P<0.05 (group C versus D); <|) P<0.05 (group A versus D); X P<0.01 (group C versus D); aa P<0.01 (group B versus D). 71 Circulating leptin levels Serum leptin levels were measured in these animals weekly to 4 weeks post-transplantation (Figure 16). Serum leptin levels were consistently higher throughout the 4 weeks of leptin measurements in group B, which received the highest volume of encapsulated leptin-producing gut cells. In week 1 post-transplantation, groups A, B, C and D had 0.3±0.1 ng/mL, 0.8±0.2 ng/mL, 0.6±0.1 ng/mL and 0.2±0.1 ng/mL leptin, respectively (P<0.01, comparing group B to group D). Week 2 post-transplantation, leptin levels were 0.7±0.1 ng/mL, 1.1±0.3 ng/mL, 0.7±0.2 ng/mL and 0.2±0.1 ng/mL for groups A , B, C and D, respectively (P<0.01, comparing group B to group D). By weeks 3 and 4 post-transplantation, serum leptin levels were not significantly different between all four groups (P>0.05). Serum leptin levels were measured in groups B and D from week 5 post-transplantation to the end of the study with no significant differences observed between the two groups (data not shown). Circulating insulin levels Serum insulin levels decreased as serum leptin levels increased (Figure 17). Serum insulin levels progressively increased in control group D throughout the 4 weeks post-transplantation: 6.3±1.7 ug/L (week 1 post), 6.7±1.5 ug/L (week 2 post), 14.4+2.3 ug/L (week 3 post) and 19.8±4.3 (xg/L (week 4 post). Insulin levels in all leptin-treated groups (groups A , B, and C) remained significantly decreased compared to group D (P<0.05) during this same time. By week 4 post-transplantation, all three leptin-treated groups had significantly lower serum insulin levels compared to control: 8.9±2.7 ug/L (group A; P<0.05), 6.1+1.2 ug/L (group B; P<0.01) and 5.9±1.2 ug/L (group C ; P<0.01). No significant differences were observed between the three leptin-treated groups. 72 2.00 1.75 3 1.50 I, 1-25 S 1.00 I 0.75 3 0.50 0.25 0.00 •P B. 2. 1. 3 1 | L c ii 0. 0. 0. 00 75-50-25-00 75 50 25 00 I I J L s c. 2.00 1.75 1.50-1 1.25 1.00 0.75H 0.50 0.25-I 0.00 • 2. 1. J 1. it £ i . C 3 o. 0. 0. 1_JL 1 • J L F igure 16. The effect of leptin cell therapy on serum leptin levels in ob/ob mice transplanted with various volumes of encapsulated control or leptin-producing gut cells. Leptin levels were measured on (A) week 1, (B) week 2, (C) week 3 and (D) week 4 post-transplantation. Leptin concentrations plotted as mean+SEM. *P<0.05, **P<0.01 (comparing all leptin-treated groups to the control group). 73 Figure 17. The effect of leptin cell therapy on serum insulin levels in ob/ob mice transplanted with various volumes of encapsulated control or leptin-producing gut cells. Insulin levels were measured on (A) week 1, (B) week 2, (C) week 3 and (D) week 4 post-transplantation. Insulin concentrations plotted as mean + S E M . *P<0.05, **P<0.01 (comparing leptin-treated groups to the control group). 74 111.4: Transplantation of reduced volumes of encapsulated leptin-producing gut cells into the intraperitoneal cavity of ob/ob mice also administered a constant dose ofRU486 (Pilot study) It was hypothesized that it could still be possible to elicit a variable degree of body weight loss in ob/ob mice by further reducing the number of transplanted encapsulated leptin-producing gut cells (to as little as 250 uL), thereby further limiting the amount of leptin being secreted. In this pilot study, on the day of transplantation, pre-transplant body weights for groups A, B and C were 63.8 g, 62.2 g and 62.5 g, respectively (Figure 18). By day 22 post-transplantation, body weight was maximally reduced to 47.8±2.6 g in ob/ob mice from group B, which received 2.5 mL of encapsulated leptin-producing gut cells. At the same time, body weight was 52.8±1.7 g in the ob/ob mice from group C, which received 250 uL of leptin-producing gut cells. Thus, mice that received the lowest volume (250 uL) of encapsulated leptin-producing gut cells lost a maximum of 9.7 g of body weight while group B, receiving a greater volume of encapsulated leptin-producing gut cells, lost more weight, 14.4 g. Mice in the control group continued to gain weight throughout the study. 75 85 80 75j 704 Transplant RU486 pellet 1 A: control cells + 87.5 ug RU486 pellet B: leptin cells (2.5mL) + 87.5 ug RU486 pellet C: leptin cells (250uL) + 87.5 ug RU486 pellet r 70 80 Duration (days) Figure 18. The effect of leptin cell therapy on body weight in ob/ob mice. Mice were transplanted with either 2.5 mL of control gut cells (group A , orange • n=1), 2.5 mL of leptin-producing gut cells (group B, blue • , n=2) or 250 uL of leptin-producing gut cells (group C, green T , n=2). Al l groups received an 87.5 pg RU486 pellet. 76 III.5: Transplantation of 500 fd. of encapsulated leptin-producing gut cells into the intraperitoneal cavity of young ob/ob mice also receiving various doses ofRU486 Body weight, food intake and blood glucose The previous pilot study (111.4) showed that varied degrees of weight loss resulted when ob/ob mice were treated with 250 uL or with 2.5 mL of encapsulated leptin-producing gut cells. To see whether transplanting a lower volume of leptin-producing gut cells with various doses of RU486 would result in RU486 dose-dependent decreases in body weight, food intake and blood glucose levels, we transplanted 500 uL of encapsulated leptin-producing gut cells or control gut cells into the intraperitoneal cavity of young ob/ob mice with various doses of RU486 (Figure 19). At the time of surgery, body weights were 43.9±1.3 g (group A, control), 43.9±1.4 g (group B, leptin-treated + 87.5 ug RU486), and 44.0±1 g (group C, leptin-treated + 8.75 ug RU486). Two weeks after transplantation body weights were 49.5±1.2 g (group A), 42.1±1.5 g (group B) and 43.8±1.2 g (group C) where control group A gained weight at a significantly higher rate than that of the leptin-treated groups B (87.5 ug RU486) and C (8.75 ug RU486) (PO.001) ; groups B and C were not significantly different from one another at this same time. From day 14 post-transplantation to the end of the study body weight gain was not significantly different between groups A, B and C (0.27±0.04 g/day, 0.32+0.05 g/day and 0.33±0.03 g/day, respectively). Food consumption also decreased in the 2 weeks after transplantation (Figure 19). At the time of transplant, food intake for groups A, B and C was 6.3+0.1 g, 6.2+0.2 g and 6.3+0.2 g, respectively, while by 2 weeks post-transplant, food intake values were 6.2±0.1 g, 3.1±0.2 g and 3.6±0.2 g respectively ( P O . 0 0 1 , comparing group A to groups 77 B and C) . Group B, which received leptin-producing gut cells and the highest dose of RU486, ate the least amount of food. Significant decreases in food intake (P<0.05) persisted in the leptin-treated groups B and C up to 4 weeks post-transplantation when compared to the control group. No significant differences in food consumption were observed between the two leptin-treated groups (B and C) throughout this time. At 5 weeks post-transplantation, group B continued to have significantly lower food intake (4.2±0.1 g) compared to group A and group C (5.0+0.03 g and 4.7± 0.1 g, respectively; P<0.01). Food intake values were no longer significantly different after 5 weeks. Blood glucose data followed a similar trend (Figure 19). On the day of transplantation, pre-transplant blood glucose concentrations were 23.1±3.5 mM (group A), 22.1±2.9 mM (group B) and 22.3±2.6 mM (group C). Two weeks following transplantation the blood glucose concentrations were as follows: group A, 26.0+2.7 mM, group B, 8.7± 0.3 mM, and group C, 10.4±0.6 mM. No significant differences were noted between the two leptin-treated groups; however, a significant difference (P<0.001) was calculated when comparing the control group A to the 2 leptin-treated groups (B and C) at this same time point. These significant differences in blood glucose concentrations in the groups B and C persisted up to day 27 post-transplantation. 78 ! SO 'Q> S 45 Transplant RU486 pellet 1 R u 4 8 6 p e | | e , 2 i ' A: control cells • 87.5 nO RU486 - B leptin cells * 87 5 ug RU486 - C : leptin calls • 8.75 mg RU486 20 B. 8 7A S 6 o Transplant RU4B6 pellet 1 30 Duration (days) RU486 pellet 2 40 3 4 5 6 Duration (weeks) 40-i 30 H E . s (J O 1 CO 20-10 Transplant RU486 pellet 1 10 50 60 Duration (days) Figure 19. The effect of leptin cell therapy with different doses of RU486 on (A) body weight, (B) food intake and (C) blood glucose in 4-5 week old ob/ob mice. Group A (orange • , n=6): 500 uL control gut cells + 87.5 jig RU486 pellets; group B (blue • , n=6): 500 uL leptin-producing gut cells + 87.5 ug RU486 pellets; group C (green A , n=6): 500 uL leptin-producing gut cells + 8.75 ug RU486 pellets . (A) Body weight, (B) food intake and (C) blood glucose values plotted as mean ± SEM. *P<0.05, **P<0.01, "*P<0.001 (comparing groups B and C to A); cP<0.01 (comparing group B to A). 79 Circulating human leptin levels Two weeks after transplantation, group A had circulating human leptin levels of 0.4±0.2 ng/mL; this was considered "background" (Figure 20). Group C (leptin-treated + 8.75 ug RU486) had the highest level of leptin, 2.95±0.24 ng/mL, and group B (leptin-treated + 87.5 ug RU486) had leptin levels of 1.78±0.85 ng/mL. It was unexpected that the treatment group receiving the lower dose of RU486 pellet (group C) appeared to produce and secrete more leptin than the treatment group receiving higher dose of RU486 (group B). This trend, however, continued to 3 weeks after transplantation, when serum leptin levels were 1.53+0.12 ng/mL in group B and 2.25±0.2 ng/mL in group C (vs. control Group A, 1.34±0.3ng/mL). Circulating insulin levels Insulin was then measured from serum samples collected prior to transplantation and once a week for 3 weeks after transplantation (Figure 21). Insulin levels in group A were 11.74±3.2 ug/L and 14.2±2.5 ug/L for the first and second weeks prior to transplant. In the first two weeks before transplantation, treatment group A had 12.47±2.3 ug/L (week 1) and 15.4+2.1 ug/L (week 2) and treatment group B had 10.7±4.1 ug/L (week 1) and 12.7±0.8 ug/L (week 2) of serum insulin. Insulin levels increased in all groups during these first 2 pre-transplant weeks. In week 1 post-transplantation, group A had an expected post-surgical decrease in serum insulin levels to 11.9±3.0 ng/mL while groups B and C had decreased insulin levels (to 9.0±1.8 ng/mL and 7.2±1.3 ug/L respectively). Although no statistical difference was found between these groups, the leptin-treated groups (B and C) had a greater decrease in insulin levels versus the control group (A) 1 week after surgery. At 2 weeks post-transplant 80 group A s insulin levels remained at 11.3±1.9 ug/L whereas the insulin levels of groups B and C continued to decrease to 8.3+2.5 ug/L and 5.5±0.9 ug/L respectively. By week 3 post-transplantation all three groups were hyperinsulinemic: group A=14.4±4.1 ug/L, group B=13.4+3.1 ug/L and group C=15.2±1.4 ug/L. 81 4n 3H ^ 2H a - J H 0 r Group A Group B Group C B. O) £ 2 Q. —i H Group A Group B Group C Figure 20. Leptin levels measured in 4-5 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of RU486. Group A: 500 uL control gut cells + 87.5 (ig RU486 pellets; group B: 500 uL leptin-producing gut cells + 87.5 ug RU486 pellets; group C: 500 uL leptin-producing gut cells + 8.75 ug RU486 pellets. Leptin levels from (A) week 2 post-transplantation and (B) week 3 post-transplantation plotted as mean + S E M . *P<0.05, **P<0.01. 82 A. 20--7 15-£ 5H 20-? 1 5 -2 3 s - 5H J l - 1 ' ' I— Group A Group B Group C Group A Group B Group C B. 20-T 1 5 -A c 10-1 "5 to £ 5-D. 20-0) i "5 V) £ 5H Group A Group B Group C Group A Group B Group C E. 20-, 3. 3 10 ^ 5H Group A Group B Group C F igu re 21 . Insulin levels measured in 4-5 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of R U 4 8 6 . Group A : 500 uL control gut cells + 87.5 ug R U 4 8 6 pellets; group B: 500 uL leptin-producing gut cells + 87.5 ug R U 4 8 6 pellets; group C : 500 uL leptin-producing gut cells + 8.75 ug RU486 pellets. Insulin levels from (A) week 1 pre-transplantation, (B) week 2 pre-transplantation, (C) week 1 post-transplantation, (D) week 2 post-transplantation, (E) week 3 post-transplantation, all plotted as mean + S E M . 83 111.6: Transplantation of 500fjL of encapsulated leptin-producing gut cells into the intraperitoneal cavity of old ob/ob mice receiving various doses ofRU486 Body weight, food intake and blood glucose Older ob/ob mice, aged 10 weeks at the start of the study, were placed in three different groups. The average body weight of each group prior to transplantation of either the control gut cells or leptin-producing gut cells was 61.0±1.2 g (group A, control gut cells + 87.5 pg RU486), 60.6±0.8 g (group B, leptin-producing gut cells + 87.5 pg RU486) and 61.2±0.8 g (group C, leptin-producing gut cells + 8.75 pg RU486) (Figure 22). Three weeks after transplantation, the control group's body weight (group A) had increased to 64.7+1.5 g, group B's body weight had decreased to 49.4±2.1 g and group C s body weight had decreased to 55.3±1.5 g. At this time, the body weight of group B was significantly different from that of both control group A (P<0.001) and group C (p<0.05). As well, group C was significantly different from control group A (P<0.01) which confirms that the two leptin-treated groups (B and C) lost weight in an RU486 dose-dependent response. In week 4 it was difficult to make objective observations regarding differences in body weight because all groups of mice were accidentally starved on days 37 and 38. However, body weights had returned to pre-starvation levels by week 5 post-transplantation. Significant differences in body weight between the 3 groups continued up to day 75 post-transplantation. Rate of body weight gain, from week 5 post-transplantation to the end of the study, was 0.2 g/day (group A), 0.4 g/day (group B) and 0.3 g/day (group C). Food intake data was also collected for 101 days and this data is presented as weekly food intake (Figure 22). In the week prior to transplantation, food intake was 7.0+0.1 g (group A), 6.5±0.1 g (group B) and 6.8±0.2 g (group C). Food intake 84 decreased significantly (PO.001) in the 2 leptin-treated groups (group B, 87.5 ug RU486 and group C, 8.75 ug RU486) compared with the control group (A) within the first several days post-transplantation. Food intake in week 1 post-transplantation was 4.6±0.5 g (group A), 2.8±0.2 g (group B), and 2.6±0.1 g (group C). In week 2 post-transplantation, food intake was significantly different between the two leptin-treated groups (B and C) versus control group A (P<0.001): 4.5+0.1 g (group A), 2.1±0.2 g (group B) and 2.9±0.03 g (group C). In week 3 post-transplantation food intake was also statistically different between the leptin-treated groups (B and C) versus control group A (P<0.001): 4.4±0.1 g (group A), 2.7±0.2 g (group B, 87.5 ug RU486) and 3.5±0.2 g (group C, 8.75 ug RU486). Food consumption by group B was significantly less than that of group C at this same time (P<0.05), again indicative of food intake reductions occurring in an RU486 dose-dependent manner. Food intake data 4 weeks post-transplantation was not reliable as all groups of mice were accidentally starved on days 37 and 38. At week 5 post-transplantation food intake for group A was 4.7±0.2 g compared to 4.6±0.2 g and 4.8±0.2 g for groups B and C, respectively (not significantly different). After week 5 post-transplantation the rate of food intake rapidly increased in the leptin-treated groups (groups B and C) at a significantly higher rate than that of the control group A (P<0.001). The rate of food intake was 0.08+0.07 g/day (group A), 0.70+0.07 g/day (group B) and 0.50±0.06 g/day (group C). By week 7 post-transplantation the food intake of the leptin-treated groups (B and C) had increased beyond the food intake of the control group: food intake was 5.6±0.1 g for group B, 5.4±0.2 g for group C, and 4.6±0.2 g for control group A . Control group A continued to consume statistically less food (P<0.01) at week 9 compared to the amount of food consumed by the leptin-treated groups (B and C). 85 Blood g lucose va lues were variable, but not statistically different, between the groups prior to transplantation (Figure 22). O n e day prior to transplantation, blood g lucose concentrat ion for control group A w a s 10.2±1.2 m M , for group B w a s 8.5±0.7 m M and for group C was 9.2±0.8 m M . F ive days after transplantation blood g lucose va lues were significantly different (P<0.05) when compar ing the blood g lucose va lues in the leptin-treated groups, B (8.5±0.3 mM) and C (9.7±0.5 mM), to control group A (11.6±0.9 mM). The significant difference, however, w a s due to an increase from pre-transplant levels in blood g lucose levels in control group A rather than a dec rease in the blood g lucose levels in groups B or C . Blood g lucose va lues of the leptin-treated groups remained the s a m e within the first post-transplantation week as the pre-transplant week. Ten days after transplantation significant dec reases (P<0.05) were observed in the blood g lucose levels of group B (7.6±0.4 mM) and group C (7.8+0.4 mM) compared to control group A (10.3±0.9 mM). Three days after the cessat ion of R U 4 8 6 re lease, on day 17 post-transplantation, the blood g lucose va lues of the leptin-treated groups, B and C , were 7.4±0.7 m M and 9.0±0.7 m M whereas the value for control group A w a s 12.7±1.4 m M (P<0.05). R a n g e s in blood g lucose va lues in all three groups, from day 17 post-transplantation to the end of the study, were variable and not significantly different from one another. 86 A. control cells + 87.5 HO RU486 4 J — B: leptin cells 87 5 (*g RU486 C: leptin » 8.75 ..g RU486 4.0 S O 6 0 7 0 D u r a t i o n ( d a y s ) 1 0 0 B . Transplant Day RU486 pellet 1 9-8-7-| « 1 -o-c. Transplant Day RU486 pellet 1 6 7 8 9 1 0 D u r a t i o n ( w e e k s ) Accidentally Starved 1 1 1 2 1 3 1 4 1 5 1 6 1 0 0 1 1 0 D u r a t i o n ( d a y s ) Figure 22. The effect of leptin cell therapy with different doses of RU486 on (A) body weight, (B) food intake and (C) blood glucose in 10 week old ob/ob mice. Group A (orange • , n=6): 500 ul_ control gut cells + 87.5 ug RU486 pellets; group B (blue • , n=6): 500 pL leptin-producing gut cells + 87.5 ug RU486 pellets; group C (green • , n=6): 500 \iL leptin-producing gut cells + 8.75 ug RU486 pellets . (A) Body weight, (B) food intake and (C) blood glucose values plotted as mean ± SEM. *P<0.05, **P<0.01, ***P<0.001 (groups B and C to A); a P<0.05 (group B to C); b P<0.01 (group B to C); c P<0.01 (group B to A); d P<0.05 (group B to A); e P<0.05 (group C to A); f P<0.01 (group C to A); g P<0.001 (group B to A). 87 Circulating leptin levels Human leptin levels were measured in plasma samples collected weekly until 4 weeks after transplantation (Figure 23). All leptin measurements were made prior to the accidental starvation of all animals. Leptin release was consistently higher in ob/ob mice from group B versus from group A and C for the first three weeks post-transplantation. In week 1 post-transplantation plasma leptin was 1.4±0.3 ng/mL in group B and 0.8±0.3 ng/mL in group C, while it was undetectable in control group A. In week 2 post-transplantation, leptin levels were: 2.4±0.1 ng/mL in group B, 1.2+0.4 ng/mL in group C and undetectable in control group A. By week 3 post-transplantation the RU486 pellets had evidently expired but leptin was still measurable in the leptin treated groups (B and C) . Group B continued to have the highest level of leptin, 2.0±0.6 ng/mL, while group C 's leptin levels decreased to 0.3±0.3 ng/mL. Finally, 4 weeks after transplantation, plasma human leptin levels in the leptin-treated groups were not significantly different from control group A. Circulating insulin levels Plasma insulin levels were measured throughout this study (Figure 24). Baseline insulin levels were measured for 2 weeks prior to transplantation and values for each group were not significantly different from one another. One week prior to transplant insulin levels were 29.4±5.4 ug/L, 18.9±1.9 ug/L, and 27.1±3.9 ug/L for groups A, B, and C, respectively. Two weeks prior to transplant insulin levels were 27.7±3.1 ug/L, 21.1 ±2.9 ug/L and 24.3±2.5 ug/L for groups A, B and C, respectively. One week after transplantation, insulin levels decreased significantly (P<0.001) in both groups B (7.3±1.7 ug/L) and C (9.6+1.9 ug/L) as compared to group A (22.1 ±3.5 ug/L). Insulin 88 levels for weeks 2, 3 and 4 post-transplantation were: 32.5±6.0 ug/L, 24.8+4.4 ug/L, and 18.2+3.9 ug/L for group A (control); 3.7±1.1 ug/L, 4.4±0.7 ug/L, and 8.8+1.1 ug/L for group B (leptin-treated + 87.5 ug RU486); 14.4±3.8 ug/L, 8.8+1.1 ug/L, and 10.7+1.9 ug/L for group C (leptin-treated + 8.75 ug RU486). Significant differences persisted when comparing insulin levels between group B (87.5 ug RU486) and group A (control): P<0.001 in weeks 2 and 3 post-transplantation and P<0.01 in week 4 post transplantation. Group C (8.75 ug RU486) also had significantly lower insulin levels as compared to group A (control) in weeks 2 and 3 post-transplantation: P<0.01 and P<0.001, respectively. Groups B and C were not significantly different from each other for the duration of 4 weeks post-transplantation. Circulating triglyceride levels Plasma triglyceride levels were also measured before and after transplantation (Figure 25). No significant differences in plasma triglyceride levels were found between the three groups within the first 2 weeks prior to transplantation or within the 4 weeks post-transplantation. In the week before transplantation, triglyceride levels were 82.1±7.7 mg/dL (group A), 76.5±6.0 mg/dL (group B) and 68.7+15.4 mg/dL (group C) . These differences were not statistically different from one another. In week 1 post-transplantation, triglyceride levels were measured at 54.3±7.0 mg/dL (group A), 57.0+5.8 mg/dL (group B) and 65.3±2.2 mg/dL (group C); again, not statistically different. All groups had an approximate 20 to 30 mg/dL decrease in pre- versus post-transplant triglyceride levels. These decreases may have been due to surgical stress since all post-transplant triglyceride levels rebounded to the pre-transplant levels by the 4th week after transplantation. 89 B Group A Group B Group C Group A Group B Group C 1 Group A Group B Group C 3.0-_2.5-E 2.0-1 Q-1.0-0) 0.5 O.O- .I Group A Group B Group C Figure 23. Leptin levels from 10 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of RU486. Group A: 500 uL control gut cells + 87.5 ug RU486 pellets; group B: 500 uL leptin-producing gut cells + 87.5 ug RU486 pellets; group C: 500 uL leptin-producing gut cells + 8.75 pg RU486 pellets . Leptin levels measured at (A) week 1, (B) week 2, (C) week 3, and (D) week 4 post-transplantation. All values plotted as mean + SEM. **P<0.01, ***P<0.001 (comparing group A to groups B and C). 90 B. Group A -i • 1 1-Group B Group C Group A Group B Group C c. D. 40-i 35-3 30-s 25-c 20-c 3 15-(0 C 10-5-0-Group A Group B Group C 40-, 35-3 30-E 25-"5> c c 20-3 15-<0 c 10-5-0- I • • 1 Group A Group B Group C E. 40-. 35-3 30-g/m 25-_c 20-c "5 15-c 10-5-0-Group A —i i— Group B Group C 40 35 3 30 t 25 ^-20 c 3 15 (0 £ 10 5' 0 Group A Group C Group B Figure 24. Insulin levels from 10 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of RU486. Group A: 500 uL control gut cells + 87.5 ug RU486 pellets; group B: 500 uL leptin-producing gut cells + 87.5 ug RU486 pellets; group C: 500 uL leptin-producing gut cells + 8.75 pg RU486 pellets . Insulin levels measured at weeks 1 and 2 pre-transplantation (A, B) , and at weeks 1, 2, 3, and 4 post-transplantation (C, D, E, and F). All values plotted as mean + S E M . **P<0.01, ***P<0.001 (comparing group A to groups B and C) . A. 125 j 100 D) E. 75 O H 50. 3 P 25 0 Group A Group B Group C 125-i 3 100-TJ a E, 75-O r - 50-s O t- 25-0-91 B. 125-. ^100-| TJ "B. g 75-O r - 50-3 £ 2SA D. Group A Group B Group C 125-1 3 100-TJ cn E, 75-(5 r - 50-3 o ( - 25-0-Group A Group B Group C Group A Group B Group C E. 125 moo TJ E 75 O 3 o 50 25 0 Group A Group B Group C 125 ••100-1 TJ O) E 75 O H 50. 3 £ 25 Group A Group B Group C Figure 25. Triglyceride levels from 10 week old ob/ob mice transplanted with a constant volume of control or leptin-producing gut cells and variable doses of RU486 Group A: 500 pL control gut cells + 87.5 pg RU486 pellets; group B: 500 pL leptin-producing gut cells + 87.5 pg RU486 pellets; group C: 500 pL leptin-producing gut cells + 8.75 pg RU486 pellets . Triglyceride levels measured at weeks 1 and 2 pre-transplantation (A, B), and at weeks 1, 2, 3, and 4 post-transplantation (C, D, E, and F). All values plotted as mean + S E M . 92 111.7: Longterm evaluation of leptin release from encapsulated leptin-producing gut cells transplanted into the intraperitoneal cavity of ob/ob mice receiving RU486 pellets Body weight, food intake and blood glucose A study was conducted to determine the duration of leptin release in vivo from the leptin-producing gut cells in the presence of RU486 (Figure 26). One day prior to transplantation, body weights for groups A, B and C were 42.1±1.6 g, 42.7±1.2 g and 42.8±1.3 g, respectively. At the end of week 1 post-transplantation the body weights of control groups A and B were 46.9±1.6 g and 44.5±1.2 g (P>0.05), respectively while the body weight of the leptin-treated group C had decreased significantly to 39.4±0.9 g (P<0.001). By week 2 post-transplantation, control groups A and B weighed 50.9±1.4 g and 44.9±1.2 g, respectively, while the body weight of the leptin-treated group C had further decreased to 37.1 ±0.9 g (PO.001) . In order to provide a constant release of RU486 to the circulation, new RU486 pellets (87.5 ug) were implanted in groups B and C every 14 days from day 10 to day 53 of the study. It was expected that cells would continue to produce and secrete leptin in the presence of RU486, thereby causing further decreases in body weight in the leptin-treated ob/ob mice group, C . On day 15, one day after the implantation of the second pellet, however, body weights of the ob/ob mice in group C started to increase and no further decreases were noted in their body weight. Rates of body weight gain, from day 15 post-transplantation to the end of the study, were 0.29±0.01 g/day (group A), 0.36±0.02 g/day (group B) and 0.51 ±0.01 g/day (group C). The rates of body weight gain were significantly different (PO.001 ) when comparing group B and C to A and comparing group B to C . 93 Food intake was calculated as weekly food consumption for each group of animals (Figure 26). One week prior to transplantation, food intake values were 6.8±1.3 g/day (group A), 6.9±1.4 g/day (group B), and 6.6±1.3 g/day (group C) . One week post-transplantation, food intake values for each group were not significantly different from one another (P>0.05): 6.9±1.0 g/day (group A), 6.1+1.3 g/day (group B) and 6.2±1.2 g/day (group C) . In week 2 post-transplantation, leptin-treated group C consumed 2.8±0.3 g/day of food, which was significantly less than the amount of food consumed by group A (6.4±0.3 g/day; P O . 0 0 1 ) and by group B (4.5±0.3 g/day; P O . 0 1 ) . After the implantation of the second RU486 pellet 3 weeks post-transplantation, group C consumed 2.2±0.3 g/day of food, which was significantly less than the food consumed by group A (6.4±0.4 g/day; P O . 0 0 1 ) or by group B (3.3±0.1 g/day; P<0.05). The decrease in food intake observed in group B was attributed to stress related to implantation of the second RU486 pellet, which required the ob/ob mice to be anesthetized during the procedure. By week 7 post-transplantation, group A consumed 5.4±0.1 g/day of food, group B consumed 4.9+0.1 g/day of food and group C consumed 6.0±0.1 g/day of food. Group C, the leptin-treated group, was consuming significantly more food than groups A (PO.05 ) and B (PO.01 ) at this time. Blood glucose levels, one day prior to transplantation, were 15.3±1.6 mM (group A) , 18.2±2.0 mM (group B) and 21.5±3.1 mM (group C) (Figure 26). Blood glucose levels, one day after transplantation were 16.8+1.6 mM (group A), 13.8±2.0 mM (group B) and 8.9±0.6 mM (group C); group C had significantly lower blood glucose levels compared to group B (P<0.05) and control group A (P<0.01). Exactly one week post-transplantation, blood glucose levels in group C were 8.3±0.6 mM which was significantly lower than group A (15.6±1.3 mM; P<0.01) and group B (13.6+1.6 mM; 94 P O . 0 1 ) . By the end of week 2 post-transplantation, blood glucose levels in group A were 10.5±1.3 mM, which was significantly higher than both groups B (6.1±0.3 mM; P<0.01) and C (5.1 ±0.5 mM; P O . 0 0 1 ) . There was no significant difference between the group that received leptin-producing gut cells (group C) and the group that received the control gut cells (group B), P>0.05. Group C continued to have blood glucose levels that were not significantly different from group B for the rest of the 60-day study. 95 Duration (days) B. Transplant 8 7 8 5 4 3 1 o RU486 pellet 1 RU486 pellet 2 RU486 pellet 3 4 5 6 Duration (weeks) Transplant 2 5 n 20 15 5H RU486 pellet 1 RU486 , pellet 2 RU486 pellet 3 , i I I 10 20 30 40 Duration (days) 50 60 70 Figure 26. The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose in 06/06 mice implanted with 3 RU486 pellets. Group A (grey • , n=6): no intervention; group B (orange • , n=6): 2 mL control gut cells + 87.5 pg RU486 pellets (x3); group C (blue W, n=6): 2 mL leptin-producing gut cells + 87.5 pg RU486 pellets (x3). Values plotted as mean ± S E M . *P<0.05, **P<0.01, ***P<0.001 (group C to A and B); a P O . 0 5 (group B to C); b P<0.01 (group B to C); c P<0.01 (group B to A); d P<0.05 (group B to A); e P<0.05 (group C to A); f P<0.01 (group C to A); h P<0.001 (group C to A); i P<0.001 (group B to A). 96 Circulating human leptin levels Leptin levels were measured from serum samples collected in the second and third week post-transplantation (Figure 27). In week 2 post-transplantation, group A (control) had undetectable serum leptin while groups B (control gut cells) and C (leptin-producing gut cells) had leptin levels of 0.2+0.1 ng/mL and 0.7±0.2 ng/mL respectively (P<0.01 when comparing group B to C) . In week 3 post-transplantation, serum leptin levels were 0.6±0.1 ng/mL (group A), 1.4+0.1 ng/mL (group B) and 3.2±0.6 ng/mL (group C). Group C continued to have significantly higher serum leptin levels than groups A (PO.001 ) and B ( P O . 0 1 ) throughout the rest of the study. Circulating triglyceride levels Triglyceride levels were measured for each group from serum samples taken at various time points: weeks 1 and 2 prior to transplantation and weeks 1, 2, 3, 5 and 6 post-transplantation (Figure 28). No significant differences were observed between any of the groups except at week 5 when group B and group C had significantly lower triglyceride levels (70.7±3.3 mg/dL and 74.1 ±8.5 mg/dL, respectively) compared with group A (97.6±4.3 mg/dL; P O . 0 1 ) . 97 A. 4 T 3 4 £ 2 H Q. —j H Group A Group B Group C B. Q. O 4n j 3 H O) ^ 2H H Group A Group B Group C Figure 27. The effect of leptin cell therapy on leptin levels in ob/ob mice implanted with 3 R U 4 8 6 pellets. Group A: no intervention; group B: 2 mL control gut cel ls + 87.5 ug R U 4 8 6 pellets (x3); group C : 2 mL leptin-producing gut cel ls + 87.5 ug R U 4 8 6 pellets (x3). Leptin levels measured at (A) week 2 and (B) week 3 post-transplantation. Al l va lues plotted as mean + S E M . **P<0.01, ***P<0.001 (comparing control group A to groups B and C ) . 98 B. 250 rr 200 | l 5 0 o I-5 o 100 so-0-250 3 200 f 150 (3 3 o G r o u p A G r o u p B Group C 100-50-0-G r o u p A G r o u p B G r o u p C 250 -i j 200 | l 5 0 O t 100H s ° 50 X G r o u p A G r o u p B Group C 250 S 200 ! E 150 100 50 0 X G r o u p A G r o u p B G r o u p C 250-3-200 5 £ 1 5 0 -100 50 0-G r o u p A G r o u p B Group C 250 rr 200 •o •& £ 1 5 0 -100 50 0 G r o u p A Group B G r o u p C 250 n rr2ooi •o E 1 5 0 H a o 100-50-0 G r o u p A G r o u p B Group C Figure 28. The effect of leptin cell therapy on triglyceride levels in oo/od mice implanted with 3 RU486 pellets. Group A: no intervention; group B: 2 mL control gut cells + 87.5 pg RU486 pellets (x3); group C: 2 mL leptin-producing gut cells + 87.5 pg RU486 pellets (x3). Triglyceride levels measured at weeks 1 and 2 pre-transplantation (A, B), and at weeks 1, 2, 3, 5 and 6 post-transplantation (C, D, E, F and G). All values plotted as mean + S E M . **P<0.01 (comparing control group A to groups B and C). 99 111.8: Longterm evaluation of leptin release from cultured encapsulated leptin-producing gut cells. This in vitro study examined the ability of encapsulated leptin-producing gut cells to produce and secrete leptin over a 5-week time frame, while in culture. Leptin-producing gut cells were grown in culture, encapsulated and cultured again for 5 weeks. Photographs of the encapsulated cells in culture were taken every week for 5 weeks (Figure 29). The encapsulated cells grew within the capsules throughout the 5 weeks. Leptin release studies were also conducted at weeks 1, 3 and 5 post-encapsulation, where cells were treated with 10"8 M RU486 then media was collected and assayed for leptin (Figure 29). At week 1, cells induced with RU486 released 270+85 ng/mL of leptin (control levels were 2.3±0.3 ng/mL). Three weeks after encapsulation the cells released 14.7±1.3 ng/mL of leptin and 5 weeks after encapsulation the cells released even less leptin, 12.8±1.1ng/mL (control leptin levels were undetectable in the absence of RU486). 100 A. 9J O » „ c. F. 300 100 j 20 T I c - I 1 n / / / / / / f j f j / j r 1 ^» ^ ° Figure 29. Evaluation of leptin release from encapsulated leptin-producing gut cells maintained in culture for 5 weeks. Photographs of encapsulated leptin-producing gut cells from week 1 (A), week 2 (B), week 3 (C), week 4 (D) and week 5 (E) post-encapsulation were incubated with 10" 8 M RU486 for 24 h and leptin was measured by ELISA (F). 101 STUDY IV: TREATMENT OF OBESITY AND DIABETES IN HIGH-FAT FED C57BL/6J MICE BY TRANSPLANTATION OF ENCAPSULATED LEPTIN-PRODUCING GUT CELLS IV.1; Transplantation of encapsulated leptin-producing gut cells into the intraperitoneal cavity of young C57BU6J mice on a high fat diet Body weight, food intake and blood glucose The potential for leptin to prevent the onset of obesity and diabetes was evaluated by administering encapsulated leptin-producing gut cells to young, lean mice prior to being placed on a high fat diet (HFD; Figure 30). Initial body weight for each group, 2 days prior to transplantation, was 23.3±0.6 g (group A , HFD, no intervention), 23.7±0.6 g (group B, regular chow (LFD), no intervention), 23.9+0.6 g (group C, HFD, control gut cells, 87.5 ug RU486) and 23.7±0.6 g (group D, HFD, leptin-producing gut cells, 87.5 ug RU486). By 3 days post-transplantation, groups C and D had lost 0.2 g and 0.3 g of body weight, respectively, while the 2 control groups, A and B, gained 0.8 g and 0.3 g, respectively. Five days post-transplantation, however, body weights were 24.3+0.6 g (group A), 24.2±0.6 g (group B), 24.9±0.7 g (group C) and 24.4±0.6 g (group D). These data were not statistically different from one another and no significant differences in body weight were observed throughout the rest of the study. Food intake was monitored on a weekly basis for the duration of this study (Figure 30 B). All four groups of mice were consuming a regular mouse chow diet prior to transplantation and the three high-fat fed groups were given the H F D beginning on the day of surgery. One week prior to transplantation, food intake was 3.0+0.1 g (group A), 3.2±0.2 g (group B), 3.0±0.1 g (group C) and 3.0±0.1 g (group D). Food consumption appeared to decrease in the high-fat fed mouse groups since 1 week after 102 transplantation, food intake for each group was 2.9±0.2 g (group A), 3.1+0.1 g (group B), 2.6±0.6 g (group C), 2.6+0.6 g (group D), but these values were not significantly different from one another. By week 5 post-transplantation, food intake for the high-fat fed groups was 2.2±0.1 g (group A), 2.2±0.1 g (group C) and 2.2+0.0 g (group D) compared with the food intake of the low-fat fed group (B), 3.1±0.0 g. Although it appeared that the high-fat fed groups of mice consumed less food, they actually were consuming 12.2 kcal (in 2 g of the 58% HFD) versus 11.6 kcal in 3 g of the low fat diet (LFD; 11% calories from fat). Blood glucose was monitored twice weekly for the duration of the study (Figure 30). Blood glucose values throughout the study were inconsistent and variable. Four days prior to transplantation, blood glucose values were 10.5+0.6 mM, 10.2+0.2 mM, 11.9±0.6 mM and 10.7+0.6 mM for group A (HFD, no intervention), group B (LFD, no intervention), group C (HFD, control gut cells) and D (HFD, leptin-producing gut cells), respectively. Three days post-transplantation, the blood glucose levels for groups A and B (10.1 ±0.8 mM and 9.8±0.5 mM) were significantly higher than those of groups C and D (7.1±0.5 mM and 8.0±0.4 mM; P<0.01 and P O . 0 5 , respectively). Seven days post-transplantation, the blood glucose values for groups A, B, C and D were 9.9±0.7 mM, 8.1±0.4 mM, 9.6±0.2 mM and 8.1±0.3 mM, respectively. Group B (control, LFD) and group D (leptin-producing gut cells, HFD, 87.5 pg RU486) had significantly lower blood glucose levels compared to groups A (HFD control) and C (control gut cells, HFD, 87.5 pg RU486) at this time ponit (PO.05) . . No further differences in blood glucose values were observed between the four groups from day 7 post-transplantation to the end of the study. 103 3 3 -*S ~ 2 8 -j= s 1 23H 0 Transplant Day 26 8.75(18 RU486 p a n t 1 pellet 2 87 S t ig RU486 pellet 3 - A. 58% fat diet, no intervention 8: regular chow, no intervention C control cells. 58%fat diet. RU486 pellets - D leptin cells. 58% fat diet. RU486 pellets 10 2 0 30 40 Duration (days) 50 60 70 B . 4 .0 3 . 5 - | - 5 3 0 2 i 2 . 5 1 2.0 1.5 .0 4 S Duration (weeks) Transplant Day 26 8 75|ig RU486 pellet 1 , pellet 2 10 87 5|ig RU486 pellet 3 30 40 Duration (days) Figure 30. The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose in C57BL/6J mice placed on a h i g h fat diet (HFD). Group A (light blue • , n=6): HFD, no intervention; group B (grey • , n=6): low fat diet (LFD), no intervention; group C (orange • n=6): HFD, control gut cells and 87.5 ug RU486 pellets; group D (dark blue T , n=6): HFD, leptin-producing gut cells and 87.5 u g RU486 pellets. All values plotted as mean ± SEM. *P<0.05, **P<0.01, ***P<0.001 (comparing group B to groups A, C and D); aP<0.05 (groups B and C to group A); aaP<0.01 (groups C and D to group A). 104 IV.2; Transplantation of encapsulated leptin-producing gut cells into the intraperitoneal cavity of obese C57BU6J mice on a high fat diet Body weight, food intake and blood glucose This study evaluated the role of leptin cell therapy in reversing obesity and diabetes in C57BL /6J mice maintained on a HFD for 40 days prior to transplantation (Figure 31). Pre-transplant body weights for groups A (HFD control), C (control gut cells) and D (leptin-producing gut cells) were 32.4+0.9 g, 31.9±1.0 g and 32.1+1.3 g, which were significantly different from the body weight of group B (LFD control), 28.5±0.7 g (P<0.05). Eight days post-transplantation, body weights were 33.7±1.1 g, 28.9±0.7 g, 31.8+0.7 g and 31.4±1.2 g for groups A, B, C and D, respectively, where group B (LFD control) weighed significantly less than group A (HFD control) (P<0.01).Fifteen days post-transplantation, body weight for groups A (HFD control, (34.9±1.3 g) and C (control gut cells, 33.0±0.5 g) were significantly higher than group B (29.4±0.8 g; P<0.01 and P< 0.05, respectively). No significant difference in body weight was noted when comparing the body weight of groups A, B, or C to that of group D (31.9±1.2g). Food intake was recorded as weekly food intake for each group (Figure 31). One week prior to transplantation, food intake was measured at 2.3+0.0 g (group A), 3.1+0.2 g (group B), 2.5±0.0 g (group C), and 2.6±0.1 g (group D). One week post-transplantation, food intake values were significantly different amongst all 4 groups (PO.001 ) 2.5±0.1 g, 3.2±0.1 g, 1.9±0.2 g, and 2.2±0.1 g for groups A, B, C and D, respectively. Beyond week 2 post-transplant, there were no significant differences in food consumption between the high-fat fed groups. 105 Blood glucose levels were measured for each group twice per week (Figure 31). Blood glucose values varied drastically from week to week and between groups even prior to transplantation. No statistical differences in blood glucose values were observed between the four mouse groups before or after transplantation of control gut or leptin-producing gut cells. 106 Transplant (Day 86) 40-, 3 5 3 ~ 30 25 20 15 RU486 release 4 58% High Fat Diet (3 groups) - A 58% HFD. no intervention B: LFD, no intervention C: 58% HFD, control gut cells + 87.5 ng RU486 - D: 58% HFD. leptin cells + 87.5 |ig RU486 10 20 30 40 B. SO 60 70 80 Durat ion (days) 6-1 s 4H • 1 3 2 90 100 110 120 130 Transplant (Week 12) 58% High Fat Diet (3 groups) RU486 release — I 1 1 1 1 1 1 1 I T I I I I 1 1 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 Durat ion (weeks) Transplant (Day 86) RLI486 release 9 0 1 0 0 1 1 0 1 2 0 1 3 0 Durat ion (days) Figure 31. The effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose in obese C57BL/6J mice on a high fat diet (HFD). Group A (light blue • , n=6): HFD, no intervention; group B (grey • , n=6): low fat diet (LFD), no intervention; group C (orange • . n=6): HFD, control gut cells and an RU486 pellet; group D (dark blue • , n=6): HFD, leptin-producing gut cells and an RU486 pellet. All values plotted as mean ± S E M . *P<0.05, **P<0.01, ***P<0.001 (comparing group B to groups A, C and D); a P<0.05 (group B to A). 107 Circulating human leptin levels Serum human leptin levels were measured at 2 weeks post-transplantation in the high-fat-fed animals only (groups A, C , and D) (Figure 32). Levels were undetectable in groups A and C. Group D, the leptin-treated group, had serum leptin levels of 2.8+1.3 ng/mL. Although these mice had increased circulating human leptin levels, it was evidently still not enough to override the leptin resistance in these animals. Circulating insulin levels Serum insulin measurements were made prior to transplantation and 2 weeks post-transplantation (Figure 33). One week prior to transplantation, insulin levels were 2.7±0.4 ng/mL (group A), 1.2±0.2 ng/mL (group B), 1.9±0.5 ng/mL (group C) and 1.7±0.2 ng/mL (group D). One week post-transplantation, insulin levels were 2.0+0.7 ng/mL, 0.9±0.1 ng/mL, 0.9+0.4 ng/mL and 0.8±0.1 ng/mL for groups A, B, C and D, respectively. No significant difference was noted when comparing group C , the group that received control gut cells, to group D, the group that received leptin-producing gut cells. Circulating triglyceride levels Serum triglyceride levels were measured prior to transplantation and one week post-transplantation (Figure 34; Table VI). Triglyceride levels were not statistically ' different amongst all 4 groups at any time point. 108 5 i cn c a _ l 3H i 2H H Group A Group C Group D Figure 32. The effect of leptin cell therapy on leptin levels in obese C 5 7 B L / 6 J mice on a high fat diet (HFD). Group A : HFD, no intervention; group C: H F D , control gut cells and an 87.5 ug RU486 pellet; group D: H F D , leptin-producing gut cells and an 87.5 \xg RU486 pellet. Leptin levels measured 2 weeks post-transplantation and are plotted as mean + S E M . **P<0.01 (comparing control group A to groups C and D). 3.5 n 3.0-5" 2.5 O) i 2.0-| 1 . 5 -I 1.0-0.5 0.0-I JL 109 B. Group A Group B Group C Group O 3.5-3.0-5" 2.5-3 2.0-c 1.5-(A 1.0-_C 0.5-0.0-Group A Group B Group C Group D 3.5 3.0-I 5" 2.5-I 3 2.0-j | 1 . 5 H I 1.0-1 0.5 0.0 Z Group A Group B Group C Group D 3.5-3.0-j 2.5 3; 2.0" | 1 . 5 -I 1.0-0.5-0.0 JL Group A Group B Group C Group D 3.5-3.0 j 2.5 3 2.0-| 1.5-I 1.0-0.5-0.0 J L X Group A Group B Group C Group D Figure 33. The effect of leptin cell therapy on insulin levels in obese C57BL/6J mice on a high fat diet (HFD). Group A: HFD, no intervention; group B: low fat diet (LFD), no intervention; group C: HFD, control gut cells and an 87.5 ug RU486 pellet; group D: HFD, leptin-producing gut cells and an 87.5 pg RU486 pellet. Insulin levels, measured at 1, 2 and 3 weeks pre-transplantation (A, B and C) and at 1 and 2 weeks post-transplantation (D and E), are plotted as mean + S E M . n=6 for each group; *P<0.05 (group B versus A). A. 125 IJ100-"a E 75-50 25 0 110 B. Group A Group B Group C Group D 125-| 100-TJ B) E. 75-O i- 50-3 o »- 25-0-Group A Group B Group C Group D 125-100-o> E. 75-O t- 50-2 t- 25-0-X X Group A Group B Group C Group D 125 U100 TJ "ft E. 75 O t 50-| H 25 X Group A Group B Group C Group D Figure 34. The effect of leptin cell therapy on triglyceride levels in obese C57BL/6J mice on a high fat diet (HFD). Group A: HFD, no intervention; group B: low fat diet (LFD), no intervention; group C: HFD, control gut cells and an 87.5 u.g RU486 pellet; group D: HFD, leptin-producing gut cells and an 87.5 pg RU486 pellet. Triglyceride levels, measured at 1, 2 and 3 weeks pre-transplantation (A, B and C) and at 1 week post-transplantation (D), are plotted as mean + S E M . n=6 for each group; *P<0.05 (compared to group A). 111 Table VI. Triglyceride levels measured from mice in Experiment IV.2 at various weeks throughout the study. Sample week T G levels (mg/dL) Group A B C D Week 1 pre- 29.43±3.4 49.7±7.0 41.6±5.1 51.6±5.8 transplant Week 2 pre- 46.1±10.1 56.4±6.6 61.1±6.2 63.7±10.5 transplant Week 3 pre- 86.5±20.9 87.2±7.4 82.4±14.5 79.6±15.2 transplant Week 1 post- 87.0±20.8 82.3±6.1 76.0±10.5 80.0±9.8 transplant Group A received a HFD with no intervention; group B received a LFD with no intervention; groups C and D received a HFD with control gut cells and leptin-producing gut cells, respectively. All values recorded as mean ± S E M . IV.3: Transplantation of a higher number of encapsulated leptin-producing gut cells into the intraperitoneal cavity of obese C57BU6J mice on a high fat diet (pilot study) In the previous high-fat fed studies (IV. 1 and IV.2), C57BL /6J mice received 5 mL of encapsulated cells at the time of transplantation. This volume of encapsulated cells was equivalent to 4.5 million control gut or leptin-producing gut cells. A s this strategy neither prevented nor reversed obesity and/or diabetes, a pilot study was done to determine if administering 4-5 times more leptin-producing gut cells could reverse obesity in these leptin-resistant C57BL/6J high-fat fed mice (Figure 35). Body weight Body weight was tracked 3 times per week for 38 days and no significant decreases were observed between the 3 groups (Figure 35). One day prior to transplantation, body weights for the 3 groups were 38.1 g (group A, HFD, no 112 transplantation, body weights for the 3 groups were 38.1 g (group A, HFD, no intervention), 40.5 g (group B, HFD, control gut cells, 87.5 pg RU486), and 41.3 g (group C , HFD, leptin-producing gut cells, 87.5 pg RU486). Five days post-transplantation, the body weights were 39.4 g (group A), 38.8 g (group B), and 39.9 g (group C). Both groups B and C had a reduction in their pre-transplant body weights of 1.7 g and 1.4 g, respectively. On day 10 post-transplantation, all groups of mice received an injection of RU486 (250 pg/kg). This was performed to rule out any drug delivery problems that may have occurred using the slow release RU486 pellets. On day 12 post-transplantation, body weights for groups A, B, and C were 37.7 g, 38.2 g, and 39.8 g, respectively. No change in body weight was observed in group C for the duration of the study. This pilot study, however, utilized very low numbers of animals (n=2) so a larger scale study was performed to mimic this pilot study and further elucidate the role of leptin cell therapy in C57BL/6J mice fed a HFD. 113 45-i Transplant of cells Injection of 250 ug/kg of RU486 B 40-| JZ m 1 I 35H! A: 58% HFD, no intervention - * - B: control cells, 58% HFD, 87.5 ug RU486 pellet C: leptin cells, 58% HFD, 87.5 ug RU486 pellet 30-10 20 Duration (days) 30 40 F i g u r e 35 . Effect of transplanting 18-million leptin-producing gut cells into the i.p. cavity of obese C57BL/6J mice (pilot study). Group A (grey • , high fat diet (HFD), no intervention; n=2); group B (orange • , HFD, 18-million control gut cells + 87.5 jig RU486; n=2); group C (blue • , HFD, 18-million leptin-producing gut cells + 87.5 ug RU486; n=2). 114 IV.4: Transplantation of a higher number of encapsulated leptin-producing gut cells into the intraperitoneal cavity of leptin-resistant C57BU6J mice on a high fat diet Body weight, food intake and blood glucose On the day of transplantation, pre-transplant body weights were 24.0±0.9 g (group A, HFD, no intervention), 25.1±0.7 g (group B, HFD, control gut cells, 87.5 pg RU486) and 25.8 g (group C, HFD, leptin-producing gut cells, 87.5 pg RU486), (Figure 36). Four days post-transplantation, body weights were 24.4±0.9 g (group A), 25.0±0.8 g (group B) and 25.5±0.9 g (group C). Body weights were not significantly different from one another for the duration of the study. Food intake was monitored weekly for each group (Figure 36). No significant decreases in food intake were observed amongst the three groups one week post-transplantation. In week 2 post-transplantation, group B had a significant increase in food intake from 2.37±0.04 g (week 1 post-transplantation) to 3.04±0.06 g (week 2 post-transplantation (PO.001)) . By week 4 and 5 post-transplantation, no significant differences in food consumption were observed. Blood glucose data are presented in Figure 36. Seven days prior to transplantation, the blood glucose level of group C was 9.0+0.7 mM; 2 days post transplantation, this group's blood glucose level dropped to 6.5±0.4 mM. This blood glucose level was significantly less than those of group A (11.1±0.8 mM; P O . 0 0 1 ) and group B (8.2±0.4 mM; P O . 0 5 ) . After this time, no further significant differences were observed between the 3 groups. 115 A. 30 28-26-£ 24 o> '© S 22 >. 1 20 18 16 Transplant (day 36) RU486 pellet 1 A: 58% HFD, no intervention - B control cells. 58% H F D . 87 5 |ig RU486 - C: leptin cells, 58% H F D . 87.5 |.g RU486 B . 10 3.5-, 3.0-| m J 2 . 5 . 1 2.0 1.5 20 30 40 Duration (days) Transplant (week 5) RU486 pellet 1 50 60 58% High Fat Diet c. 4 6 Duration (weeks) 10 58% Hign Fat Diet 1^ Transplant (day 36) RU486 pellet 1 . 11-| 1 0 -O 8 6 10 20 30 40 50 60 Duration (days) Figure 36. Effect of leptin cell therapy on (A) body weight, (B) food intake and (C) blood glucose in obese C57BL/6J mice after transplanting 18-million control or leptin-producing gut cells. Group A (grey • , high fat diet (HFD), no intervention; n=6); group B (orange • , HFD, 18-million control gut cells + 87.5 pg RU486; n=6); group C (blue T , HFD, 18-million leptin-producing gut cells + 87.5 pg RU486; n=6). Values plotted as mean ± S E M . ***P<0.001 (group C versus B); bP<0.01 (group C versus B); cP<0.01 (group B versus A); dP<0.05 (group B versus A); hP<0.001 116 Circulating human leptin levels Serum human leptin levels were measured in groups B and C only, at 2 weeks post-transplantation (Figure 37). Group B had undetectable circulating human leptin while group C had 12.3±3.1 ng/mL of human leptin in their serum. Even though group C had significantly more leptin in their serum than the control groups, it was not enough to overcome the apparent leptin resistance in these high fat fed obese C57BL /6J mice. Circulating insulin levels Insulin levels in these animals are shown in Figure 38. Insulin levels were not significantly different amongst groups prior to transplantation. One week post-transplantation, insulin levels for groups A, B and C were 0.7±0.1 ng/mL, 0.5±0.1 ng/mL and 0.5±0.1 ng/mL, respectively. Groups B and C were significantly less than group A (P<0.05), but not from each other. No further significant decreases were measured between the three groups in weeks 2 or 3 post-transplantation. Circulating triglyceride levels In the week prior to transplantation, serum triglyceride levels were 113.9±10.7 mg/dL (group A HFD, no intervention), 90.1±13.5 mg/dL (group B, HFD, control gut cells), and 140.8±15.0 mg/dL (group C, HFD, leptin-producing gut cells) (Figure 39). In week 1 post-transplantation, serum triglyceride levels were 99.7±9.7 mg/dL (group A), 57.9+3.8 mg/dL (group B), and 78.8±3.1 mg/dL (group C) . Group B was significantly lower than groups A (P<0.001) and C (P<0.05). In week 2 post-transplantation, serum triglyceride levels for groups A, B and C were 144.8±15.8 mg/dL, 54.1+4.0 mg/dL, and 90.7+14.8 mg/dL, respectively, where groups B (control gut cells) and C (leptin-117 producing gut cells) had significantly lower triglyceride levels than group A (HFD control, P<0.001 and P<0.01, respectively). 118 20n 15 o> S 10-c Q. O 5H ** Group B Group C Figure 37. Effect of leptin cell therapy on leptin levels in obese C 5 7 B L / 6 J mice after transplanting 18-million control or leptin-producing gut cel ls . Group B: high fat diet (HFD) , control gut cells and an 87.5 | i g R U 4 8 6 pellet; group C : H F D , leptin-producing gut cells and an 87.5 ug R U 4 8 6 pellet. Leptin levels measured 2 weeks post-transplantation and are plotted as mean + S E M . **P<0.01 (comparing group B to group C) . A. 119 B. 1.5n a. |o.5H c. 0.0-1.5 Group A Group B Group C Group A Group B Group C SS0.5H 0.0 1.5-2 0.5 Group A Group B Group C 0.0 Group A Group B Group C i 1 1 r Group A Group B Group C Figure 38. Effect of leptin cell therapy on insulin levels in obese C 5 7 B L / 6 J mice on a high fat diet (HFD), transplanted with 18-million control or leptin-producing gut cells. Group A (HFD, no intervention; n=6); group B (HFD, 18-million control gut cells + 87.5 pg RU486; n=6); group C (HFD, 18-million leptin-producing gut cells + 87.5 pg RU486; n=6). Insulin levels measured at weeks 1 and 2 pre-transplantation (A, B) and at weeks 1, 2 and 3 post-transplantation (C, D and E). Values plotted as mean + S E M . *P<0.05 (groups B and C versus A). 120 B. 200-i 3 150H E o 50H 200 rj 150-1 "3) E o 100H r-o 50H Group A Group B Group C Group A Group B Group C C. 200-i 5 150-1 E o 100-1 H 2 o 50-200-i 5 150-1 "B> E o 100H H o 50-j Group A Group B Group C Group A Group B Group C Figure 39. Effect of leptin cell therapy on triglyceride levels in obese C 5 7 B L / 6 J mice on a high fat diet (HFD), transplanted with 18-million control or leptin-producing gut cells. Group A (HFD, no intervention; n=6); group B (HFD, 18-million control gut cells + 87.5 pg RU486; n=6); group C (HFD, 18-million leptin-producing gut cells + 87.5 pg RU486; n=6). Triglyceride levels measured at weeks 1 and 2 pre-transplantation (A, B) and at weeks 1 and 2 post-transplantation (C and D). Values plotted as mean + S E M . *P<0.05 (group C versus A and B); a P<0.05 (group C versus B); f P O . 0 1 (group C versus A) ; g P O . 0 0 1 (group B versus A). 121 IV.5: Effect of inducing encapsulated leptin-producing gut cells with RU486 24-hour s prior to transplantation into obese C57BU6J mice (a pilot study) Body weight, food intake and blood glucose One week prior to transplant, all control gut cells and leptin-producing gut cells were encapsulated and maintained in culture (Figure 40). On the day before surgery, leptin-producing gut cells were treated with 10"8 M RU486. No change in body weight was observed in treated animals following transplantation of encapsulated cells; post-transplant body weights of groups A and B were not significantly different from their pre-transplant body weights. Two days prior to transplantation, the body weight for group A was 27.9 g and for group B' was 28.1 g. Five days post-transplantation, the body weight for group A was 28.1 g and for group B was 28.3 g. Food intake decreased 1 day after surgery by 0.6 g in both groups. Food intake gradually increased after this time and no significant differences were observed. Blood glucose concentrations were measured at 8.1 mM (group A) and 10.3 mM (group B) 4 days prior to transplantation. Three days post-transplantation, blood glucose levels were measured at 7.4 mM (group A) and 10.9 mM (group B); blood glucose levels in group B were not significantly altered comparing pre- to post-transplant values. 122 A. 30 29 | 28 i a 27-26 Encapsulation of control or leptin-producing cells Induction of encapsulated cells with 10"M RU486 for 24 hours A: control cells, 58% HFD, 87.5 ug RU486 - 0 leptin cells, 58% HFD, 87.5 ig RU486 B. 8 10 12 Duration (days) 14 16 18 20 s 2 S • •S 1 c o o 2.04 15 Encapsulation of control or leptin-producing cells Induction of encapsulated cells with 10'*M RU486 for 24 hours I 12 14 16 18 20 Time (days) c. 12-1 10 1 Induction of encapsulated cells with 10"*M RU486 for 24 hours Transplant 8 10 1 Time (days) Figure 40. Effect of transplanting 18-million encapslated control or leptin-producing gut cells, induced with RU486 24 h prior to transplantation, on (A) body weight, (B) food intake and (C) blood glucose in obese C57BL/6J mice (pilot study). Group A (orange • , high fat diet (HFD), control gut cells + 87.5 pg RU486); group B (blue T , HFD, leptin-producing gut cells + 87.5 pg RU486). 123 DISCUSSION Regulation of leptin mRNA and protein expression and secretion from leptin-producing gut cells Leptin is released primarily from white adipose tissue and transfers information on the energy stores of the adipose tissue to the hypothalamus (18,19). Generally, plasma leptin levels remain at a steady state within individuals and vary according to body mass index values (82). Administration of leptin by injection to treat obesity and diabetes in ob/ob mice has been effective in reducing body weight, food intake and blood glucose levels (35-38); however, leptin gene therapy has proven to be more effective, yielding more weight loss within a shorter period of time despite peak leptin levels approximately ten times less than that of animals treated by leptin injection (75, 76). The main limitation of gene and cell therapy systems is that they lack effective post-translational control of transgene expression. In order to address this problem, several research groups have developed artificial regulatable expression systems that will activate transgene expression in the presence of a drug (94, 95, 109, 110). In 1994, Wang et al. developed an inducible GeneSwitch™ System where a target gene is expressed in the presence of an exogenous stimulus, RU486 (94). In our lab, leptin-producing gut cells were created by transfecting GTC-1 cells with the modified GeneSwitch™ system plasmids encoding the human leptin protein. Our in vitro studies revealed that when these leptin-producing gut cells were induced with 10"8 M RU486, for various time points, leptin m R N A expression (Figure 4), leptin protein expression (Figure 5), and leptin protein release (Figures 6 & 13A) increased in an RU486 time-dependent manner. Additionally, when these cells were induced with various doses of RU486 for 24 h, increases in leptin mRNA, leptin protein expression and leptin protein release (Figures 4, 5, 6, 13 A) were RU486 dose-dependent. Importantly, leptin m R N A 124 expression and leptin protein expression were not evident when no RU486 was present. These data support previous findings that activation of transgenes using the RU486 inducible GeneSwitch™ does not occur unless RU486 is present (94, 96), and that it is possible to elicit an RU486 dose-dependent release of the transgene of interest (96). Since the leptin-producing gut cells were originally derived from GTC-1 cells, which normally produce and secrete GIP, it was of interest to determine whether GIP expression was altered in the leptin-producing gut cell line. GIP m R N A expression was quantified from GTC-1 cells, control gut cells and leptin-producing gut cells and minimal differences in GIP m R N A expression were observed in comparing all three gut cell lines (Figure 3). The GeneSwitch™ system has previously been shown to have no adverse or toxic effects on cells and tissues (96) and our data support this observation. Encapsulating cells in a sodium-alginate membrane or coating allows cells to be immuno-isolated while allowing nutrient influx and cell product efflux across the semi-permeable membrane (104, 111). We evaluated in vitro leptin protein secretion from encapsulated leptin-producing gut cells after being induced with various concentrations of RU486. Encapsulated leptin-producing gut cells appeared to secrete about 50% less leptin than free, non-encapsulated leptin producing gut cells (Figures 13A and B). However, the leptin protein release profile from the encapsulated leptin-producing gut cells was the same as that observed from the free, non-encapsulated leptin-producing gut cells. The decrease in leptin secretion was unexpected since other researchers have measured similar amounts of secreted protein from encapsulated versus from free cells (103), and when comparing protein release from encapsulated islets to free islets (112). In our experiments, we did not ensure that the exact same number of encapsulated versus non-encapsulated cells were being incubated in RU486 and it is possible that a variability in the number of cells may account for the leptin level 125 difference in these experiments. It has also been reported that it is possible to improve the yield of secreted recombinant proteins from encapsulated cells by optimizing the encapsulation procedure through identifying optimal bead size, cellular density, homogeneity and ion composition of the gel (104, 106). Leptin therapy by injection is short-lived due to the rapid degradation of the protein in the bloodstream (76). The duration of leptin release, therefore, is another important consideration in developing an effective leptin cell therapy. In our in vitro experiments, encapsulated leptin-producing gut cells were cultured for 5 weeks and leptin release was measured at weeks 1, 3 and 5 post-encapsulation (Figure 29). A dramatic decrease in leptin secretion from the encapsulated leptin-producing gut cells was observed over a 5-week period. Previous studies observed variations in the secretion levels of proteins from encapsulated cells and this was attributed to differences in the number of cells within each capsule (91). It could be possible that in our studies a variation in the actual number of capsules induced with RU486 may have been variable from week to week. We had measured the same volume of capsules per week but it is conceivable that there could have been variation in the number of capsules within these measured volumes. Visual examination of GTC-1 cells and leptin-producing gut cells grown in a monolayer revealed that these cells continued to grow until they were approximately 85-90% confluent, at which point they started to die. It was apparent that these same cells, when grown within a microcapsule, grew in a 3-D formation whereby they formed large cellular aggregates. These large cellular aggregates could cause cells, trapped inside of these formations, to become compromised thereby resulting in less leptin production and release. Zhou etal. (103) made similar observations with encapsulated 6TC6-F7 cells in culture. They described the BTC6-F7 cells growing into large 3-dimensional aggregate formations within the 126 capsules. Following C a 2 + imaging, Zhou et al. also confirmed that the central core of these aggregates may be less accessible to larger molecules that can pass into the capsules leading to a decrease in the responsiveness of the cells in the central core (103). Other investigators report that the greater the "dead space" within the capsules, the slower the encapsulated cell response will be (72). A s the leptin-producing gut cells in our studies formed these large cellular aggregates, cells were no longer dispersed evenly throughout the capsule creating more "dead space". All of observations of the behaviour of encapsulated leptin-producing gut cells in our studies support many of the observations made by previous investigators. Treatment of diabetes and obesity in ob/ob mice by leptin cell therapy Ob/ob mice are leptin deficient and are characterized by marked obesity, hyperphagia, transient hyperglycemia and hyperinsulinemia (34). Ob/ob mice treated with leptin respond by losing weight, consuming less food, increasing energy expenditure and lowers glycemia (35-38, 75, 76). Our experiments evaluated the efficacy of treating obesity and diabetes in ob/ob mice with a leptin cell therapy. In our kidney capsule studies (11.1 and II.2), decreases in body weight and food intake were observed following leptin therapy. Ob/ob mice that received leptin-producing gut cells to their kidney capsules, simultaneously implanted with an RU486 pellet, lost up to 7% of their body weight between days 8 and 13 post-transplantation, at the same time they consumed 63% less food per day (Figure 7). When ob/ob mice were transplanted with leptin-producing gut cells to their kidney capsule, implanted with RU486 and given C s immunosuppressant therapy, they lost up to 11% of their body weight between days 11 and 17 post-transplantation; at the same time, these ob/ob mice consumed 4 0 % less food per day. Without immunosuppressant therapy, it 127 appears that transplanted leptin-producing gut cells are less effective in releasing leptin after 13 days post-transplantation as compared to those mice that received C s . Immunosuppressant therapy appears to prolong the viability of the transplanted leptin-producing gut cells, under the kidney capsule, thereby allowing for more profound effects of leptin on body weight and food intake in ob/ob mice. In the ob/ob mice treated with encapsulated leptin-producing gut cells (Figures 14, 15, 18, 19, 22, and 26), transient reductions in body weight and food intake were observed, generally lasting no longer than 14 to 17 days post-transplantation. The length of time that the leptin-producing gut cells were able to decrease body weight and food intake were not significantly different from what was observed in the reductions in ob/ob mice that received leptin-producing gut cells to their kidney capsule, with RU486 and C s therapy. In study III.7 (Figure 26), we had attempted to cause long-term leptin release in ob/ob mice transplanted with leptin-producing gut cells in the presence of RU486. Unfortunately, body weight and food intake reductions were not prolonged beyond 14 to 17 days even though 3 RU486 pellets had been implanted at 14-day intervals from the time of transplantation. The second and third RU486 pellets were implanted subcutaneously at the base of the neck while the initial RU486 pellet was implanted into the i.p. cavity where all of the encapsulated leptin-producing gut cells were injected. It is possible that delivery of RU486 to the encapsulated leptin-producing gut cells from the second and third RU486 pellets was not as effective as the delivery from the first RU486 pellet because of the location of implantation. Alternatively, it could be that the transplanted cells do not survive longterm (14-17 days) in the peritoneal cavity due to the lack of vascularization and decreased access to nutrients (104). Another explanation for the transient body weight and food intake decreases could be due to the fact that after 2 weeks of being in the ob/ob mice, the encapsulated leptin-producing gut 128 cells may have been killed by cytokines being released from immune cells within the ob/ob mice. This explanation is not as likely since other studies have shown that encapsulated cells remain viable between 60 days to 24 weeks in vivo (91, 103). It is also possible, in considering the results from the long-term in vitro study (III.8; Figure 29), that over time the leptin-producing gut cells have formed large cellular aggregates preventing RU486 from stimulating the cells in the central core to produce and secrete leptin. This, in turn, may explain why reductions in body weight or food intake were not prolonged after 2 weeks post-transplantation. Our data support observations made by previous studies of ob/ob mice treated with leptin (35-38, 75, 76). Researchers observed that when ob/ob mice received leptin treatment, not only did the mice lose weight, consume less food and have improved glucose homeostasis, they responded in a leptin dose-dependent manner. Ob/ob mice injected with 300 pg of leptin daily had severe suppression of food intake but when the leptin dose was decreased to 100 pg of leptin, the ob/ob mice still weighed less than the controls but were not starving or malnourished (38). Similar to our findings, Pelleymounter et al. observed a 22% decrease in body weight values and a 52.6% reduction in food consumption in ob/ob mice injected with 10 mg/kg of leptin, daily over a 28-day study (37). Miniosmotic pump studies revealed a 30-40% reduction in body fat mass when continuously administering 10 pg/day of leptin (113). Transplanting as little as 450,000 leptin-producing gut cells (as in our studies) results in leptin levels that are similar to those observed in mice receiving daily injections of 10 mg/kg of leptin and in mice receiving continuous infusions of 10 pg/day of leptin by miniosmotic pump. Interestingly, in the kidney capsule studies (11.1 and II.2), blood glucose concentrations were reduced by 70% beyond 30 days post-transplantation in leptin-treated ob/ob mice, regardless of whether they were treated with C s or not (control 129 ob/ob mice remained hyperglycemic). These data suggest that longterm release of low levels of leptin potently regulates blood glucose concentrations levels since post-transplant blood glucose concentrations remained normalized at the same time that post-transplant body weight and food intake values increased to pre-transplant levels. Alternatively, transient leptin production may have longterm effects on glucose homeostasis. The observations of long-term blood glucose normalization were confirmed in studies III.2, III.3, and III.5 where ob/ob mice were transplanted with encapsulated leptin-producing gut cells to their i.p. cavity in the presence of RU486 (Figures 14 ,15, 19). Study III.2 (Figure 14) revealed that blood glucose concentrations remained reduced by 70% for up to 50 days post-transplantation, in leptin-treated ob/ob mice. In study 111.3 (Figure 15), all three groups of ob/ob mice that were transplanted with encapsulated leptin-producing gut cells had blood glucose concentration reductions of up to 65-70% that lasted beyond 50 days post-transplantation. Study III.5 (Figure 19) showed similar decreases in the leptin-treated ob/ob mice - reductions of up to 64% in blood glucose levels were observed lasting beyond 30 days post-transplantation. Remarkably, in all of these studies, blood glucose normalizations persisted even when body weight and food intake in the leptin-treated ob/ob mice had increased beyond pre-transplant values. Pelleymounter er al. observed that administering 0.1 mg/kg of leptin by injection did not promote body weight loss but did decrease blood glucose and insulin levels (37). Reductions in blood glucose levels without changes in body weight were also observed after treating ob/ob mice with leptin gene therapy (75, 76). Furthermore, when a single intraperitoneal injection of leptin was administered to ob/ob mice, reductions in blood glucose levels were observed after 24 h, independent of any changes in body weight (77, 78). All of these data suggest that leptin is a potent regulator of blood glucose and can have direct effects on blood glucose and insulin 130 levels, independent of influences on body weight and food intake. Our leptin cell therapy provides long-term blood glucose normalization even when body weight and food intake values have returned to pre-transplant values. This leptin cell therapy may cause prolonged normalization of blood glucose levels if the cells are "leaky", leading to small increases in circulating leptin levels that may be undetectable by leptin RIA. However, it is unlikely for these leptin-producing gut cells to secrete leptin in the absence of RU486 since our in vitro studies confirmed that leptin was released from these cells only in the presence of RU486. Alternatively, it could be proposed that continuous delivery of leptin for 14 days by leptin cell therapy has potentially long-lasting effects on insulin sensitivity by altering gene expression of target genes. This possibility warrants further investigation. It is important to point out that in studies 111.3 and III.5, when the ob/ob mice in the control groups reached approximately 9-10 weeks of age, they also became normoglycemic. In these 2 encapsulation studies (III.3 and III.5), we cannot conclude that the small doses of leptin secreted from the encapsulated leptin-producing gut cells were definitively the cause of normoglycemia in the leptin-treated ob/ob mice since no significant difference in blood glucose levels were noted as compared with the control ob/ob mice. Our data show that hyperglycemia in ob/ob mice is variable and sometimes transient. Although the transient hyperglycemia in ob/ob mice has not been scientifically explained, other researchers have documented that once ob/ob mice reach a certain age they revert to a normoglycemic state (34). Reductions in body weight were observed in the group of ob/ob mice that were transplanted with control gut cells to their kidney capsule (Figure 9, study II.2). In this ob/ob kidney capsule experiment, the 2 groups of mice treated with C s both lost body weight. The ob/ob mice that received leptin-producing gut cells, RU486 and C s lost 131 weight within the first 3 weeks post-transplantation, whereas the ob/ob mice that received control gut cells, RU486 and C s started to lose weight after the third week post-transplantation. The leptin producing gut cell-treated group started regaining their body weight within the fourth week post-transplantation but the control gut cell-treated group continued to gradually lose weight to the end of the study. It is important to point out that our data do not support the hypothesis that C s therapy was what was causing the body weight reductions in these mice. The physical appearance of these two groups of ob/ob mice was drastically different from each other. The leptin-treated ob/ob mice appeared to be better groomed with much healthier skin integrity compared to the control ob/ob mice (Figure 11). Most importantly, the blood glucose levels of the leptin-treated ob/ob mice were normalized for the majority of the study whereas the control ob/ob mice remained hyperglycemic for the duration of the study (Figure 9). This suggests that it was the diabetes that caused these mice to lose weight by the end of the study and not the C s treatment since the group of mice that received leptin and C s had a more healthy appearance. These observations are supported by previous studies that examined the effect of leptin on wound healing in ob/ob mice. Leptin has been observed to improve and promote wound healing in ob/ob mice (114). Leptin has been described to be a regulatory link between the endocrine system and the immune system with respect to skin repair (115). Ob/ob mice lose weight and reduce their food intake in a leptin dose-dependent manner (35-38, 75, 76). One of the goals of our experiments was to try and elicit a dosed release of leptin from leptin-producing gut cells, in vivo, by varying the concentration of RU486 being administered to ob/ob mice. Leptin release from leptin-producing gut cells is RU486 dose-dependent in vitro (Figure 6) so we attempted to determine if leptin could be released in an RU486 dose-dependent manner in vivo, 132 thereby causing a leptin dose-dependent reduction in body weight, food intake and blood glucose levels in ob/ob mice. In study III.2 (Figure 14), 1 mL of encapsulated leptin-producing gut cells was transplanted into ob/ob mice while RU486 pellets of various doses were implanted into different groups (Figure 14). Although it was expected that the different groups of ob/ob mice would respond with variable decreases in body weight and food intake, all three groups of mice that received leptin-producing gut cells with various doses of RU486 pellets responded with the same degree of weight loss and food intake reductions. It could be possible that the amount of leptin being secreted after induction with even the lowest dose of RU486 (0.875 \ig pellet) was enough to saturate a large number of leptin receptors in the ob/ob mice thereby leading to maximal decreases in body weight, food intake and blood glucose concentrations. The next encapsulation study, study 111.3 (Figure 15), maintained constant RU486 doses while the volume of transplanted leptin-producing gut cells injected intraperitoneally was varied (Figure 15). It was expected that by decreasing the number of leptin-producing gut cells being transplanted, the smaller number of cells would release the smallest concentrations of leptin and lead to a lesser decrease in body weight and food intake compared to the other groups receiving more leptin-producing gut cells. The results, however, showed the same amount of body weight, food intake and blood glucose reductions in all leptin-treated groups, independent of how many leptin-producing gut cells were transplanted into each group. Interestingly, the measured serum leptin concentrations (Figure 16) revealed RU486 dose-dependent leptin release, even though no leptin dose-dependent decreases were observed in body weight and food intake. The higher the volume of leptin-producing gut cells transplanted, the more leptin was secreted. Serum insulin levels responded in parallel to the leptin concentrations - the more leptin measured, the lower the measured plasma 133 insulin concentrations (Figure 17). These data confirmed that it was possible to secrete variable levels of leptin via leptin cell therapy even though leptin dose-dependent decreases in body weight, food intake and blood glucose levels were not observed. These data also suggest that these mice resonded maximally to even the lowest administered dose of leptin. It is interesting to note that the percentage of body weight loss and food intake reductions were very similar in our experiments to those observed in the studies performed by Pelleymounter et al. when they administered the highest dose of leptin (10 mg/kg/day) (37). This suggested that in our experiments, the leptin receptors may have been fully saturated even at the lowest level of leptin release, which would explain why body weight and food intake reductions did not occur in a leptin dose-dependent manner. In study III.5 (Figure 19), we attempted to elicit a leptin dose-dependent decreases in body weight, food intake and blood glucose levels by drastically reducing the number of encapsulated leptin-producing gut cells being transplanted into ob/ob mice. Ob/ob mice were transplanted with 500 pL of leptin-producing gut cells with various doses of RU486. Both leptin-treated groups responded with similar reductions in body weight, food intake and blood glucose levels even though they were administered different doses of RU486. This same experimental protocol was repeated in study III.6 (Figure 22), using older ob/ob mice. Older ob/ob mice responded with RU486 dose-dependent decreases in body weight, food intake and blood glucose levels, which suggested that the variable RU486 doses caused secretion of variable levels of leptin thereby leading to leptin dose-dependent decreases in body weight, food intake and blood glucose levels. These data do support the findings that ob/ob mice lose weight and reduce their food intake in a leptin dose-dependent manner (35-38, 75, 76), and these data also prove that the leptin cell therapy system is effective in vivo. 134 Our leptin cell therapy studies have revealed different degrees of body weight loss in ob/ob mice of different ages. This indicates that the age of the ob/ob mice is critical in determining how much body weight can be lost after being transplanted with encapsulated leptin-producing gut cells in the presence of RU486. In studies III.4 and 111.6, the ob/ob mice were approximately 10 weeks of age or slightly older (Figures 18 and 22). The leptin-treated ob/ob mice in study III.4 lost a maximum of 2 3 % of their body weight and the leptin-treated ob/ob mice in study III.6 lost a maximum of 19% of their body weight. These differences were not significantly different from the body weight losses observed by others (35-38, 75, 76). Ob/ob mice that received encapsulated leptin-producing gut cells in study III.5 were 4-5 weeks old. Their body weight reductions were minimal since they lost only 4% of their body weight while the control gut cell-treated ob/ob mice gained 12% of their body weight at the same time. It is apparent that body weight gain is blunted in young ob/ob mice since body weight loss is not as profound as seen in the older ob/ob mice. It's interesting to note that younger ob/ob mice weigh less than older ob/ob mice and therefore have less potential body fat to lose. It is likely that since these older ob/ob mice have more body fat mass, their peripheral and central leptin receptors may be upregulated to a higher degree than what might be occurring in the younger ob/ob mice. Leptin receptor m R N A is upregulated in ob/ob mice (116, 117) and ob/ob mice are five-times more sensitive to leptin administration than lean controls (36). We speculate that since younger, lighter ob/ob mice appear to be less sensitive to leptin, they may have less upregulation of their leptin receptors than older, heavier ob/ob mice, which would explain the blunted response in body weight reduction in the younger ob/ob mice. In a previous leptin gene therapy study comparing leptin gene therapy to daily leptin injection therapy, Morsy et al. observed a higher rate of weight loss in the gene 135 therapy-treated group of ob/ob mice compared with the ob/ob mice receiving daily leptin injections. A s well, they observed that leptin gene therapy produced peak leptin levels approximately 10 times less than that of animals treated by leptin injection yet yielded more weight loss within a shorter period of time (76). Treatment by leptin injection requires large doses to be administered daily, as observed in previous studies (35-38). Morsy et al. injected a total of 50 ug of leptin per day and achieved a 22.9% reduction in body weight in ob/ob mice; the serum leptin levels measured on day 4, 1 h after injection in these same animals was 335.2±52.9 ng/mL (76). At the same time they had treated ob/ob mice with human leptin by adenovirus-mediated delivery and observed a 22.3% reduction in body weight; the serum leptin levels 4 days after intervention in these animals was measured to be 22.9±2.6 ng/mL. They were able to observe a similar maximum body weight loss in both the leptin gene therapy treated and leptin injected-treated animals even though the amount of circulating serum leptin was drastically different in the two groups. Our leptin cell therapy studies, either by kidney capsule transplantation or injection of encapsulated leptin-producing gut cells, support the findings by Morsy et al. whereby a small dose of circulating leptin, administered by leptin cell therapy, is very effective in treating obesity and diabetes in ob/ob mice. Our studies confirmed that it is possible to treat diabetes and obesity in ob/ob mice using a regulatable leptin cell therapy; as well, treatment of obesity and diabetes can be controlled in ob/ob mice in an RU486 dose-dependent manner. Leptin cell therapy in the treatment of obesity and diabetes in high-fat fed C57BL/6J mice The majority of obesity cases in rodent models of obesity (118) and in humans (82) are associated with high levels of circulating leptin and leptin resistance. Rodents 136 fed a high fat diet (HFD) will develop obesity associated with hyperleptinemia and will become insensitive to peripherally and centrally administered leptin (87, 88, 119). In an attempt to prevent the development of obesity and diabetes in mice fed a H F D , we treated C57BL/6J mice with leptin cell therapy prior to placing the mice on the 58% HFD. Minimal decreases in body weight, food intake and blood glucose values were observed, and these decreases were attributed to surgical stress rather than the effects of leptin. Previous findings that showed when rats were administered leptin gene therapy centrally, prior to consumption of a HFD, they did not gain weight and did not suffer from hyperinsulinemia, suggesting that leptin resistance may be caused by a leptin transport defect at the blood-brain barrier (120) Our data support the hypothesis that leptin resistance may involve the transport of leptin across the blood-brain barrier since peripherally administered leptin, by our leptin cell therapy did not prevent the onset of obesity in diet-induced obese mice. Another objective in the treatment of obesity is the reversal of obesity in high fat-fed rodents. In study IV.2, C57BL/6J mice were fed a HFD for over 80 days prior to receiving leptin cell therapy (Figure 31). C57BL/6J mice consuming a HFD in our studies gained a moderate amount of body weight but did not develop hyperinsulinemia or hyperglycemia as expected. Minimal decreases in body weight, food intake and blood glucose levels were observed in both control gut cell-treated and leptin-producing gut cell-treated C57BL /6J mice. These decreases were again attributed to surgical stress since the leptin-treated group did not respond differently from the control-treated group of mice. Human leptin levels measured in C57BL/6J mice, transplanted with leptin-producing gut cells, were 2.8±1.3 ng/mL, similar to the leptin concentrations we had measured in the ob/ob studies. This concentration of leptin was likely not enough to override leptin resistance since ob/ob mice are much more sensitive to leptin than 137 lean controls (36). Also, Van Heek et al. found that in order to decrease food intake in high fat fed C57BL /6J mice, they needed to inject 25-times more leptin than what was previously injected into ob/ob mice to reverse obesity and diabetes (87). Clearly, the amount of leptin we had administered into the mice in studies IV.1 and IV.2 was not enough to prevent the onset of obesity and diabetes and was not enough to override the leptin resistance that develops in these high fat fed mice. Interestingly, even after transplanting 18 million encapsulated leptin-producing gut cells (Study IV.3, Figure 35), 4-times the number of cells transplanted into the previous HFD study (IV.2), no changes in body weight, food intake or blood glucose levels were observed. Serum human leptin levels had also increased approximately 4-fold more than the previous H F D study but still the amount of leptin being administered by leptin cell therapy was not enough to override the leptin resistance in these high-fat fed, obese C57BL/6J mice. Our data confirm that peripheral administration of higher doses of leptin, by leptin cell therapy under the conditions tested, does not override the leptin resistance that develops in high fat fed obese C57BL/6J mice. Our data support numerous other findings that the amount of peripherally and centrally administered leptin, either by intracerebral ventricular injection, subcutaneous injection or gene therapy, has not been high enough to reverse obesity and diabetes in diet-induced obese rodents (89, 90, 116). 138 CONCLUSIONS Reversal of obesity and diabetes in ob/ob mice has previously been effective with leptin treatment either by injection or by virally mediated gene therapy. Our studies revealed that administering leptin by a leptin cell therapy is efficacious in reversing obesity and diabetes in ob/ob mice. We have concluded that leptin release from leptin-producing gut cells can be regulated by switching leptin transgene expression "on" or "off' in the presence of a drug (RU486). We also established that it is possible to control the amount or dose of leptin being secreted from leptin-producing gut cells by altering the dose of RU486 being administered. Ob/ob mice lost body weight and had reductions in food intake after transplantation of encapsulated leptin-producing gut cells, only in the presence of RU486. We provide evidence that it is possible to elicit an RU486 dose-dependent decrease in body weight and food intake in vivo, in ob/ob mice. Our studies reveal a remarkable long-term normalization of blood glucose concentrations, independent of changes in body weight and food intake, in ob/ob mice that received leptin cell therapy. 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