NOVEL ENTEROENDOCRINE RECEPTORS REGULATING SECRETION AND GLUCOSE HOMEOSTASIS

by

Grace Beatrice Flock

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Institute of Medical Science, University of Toronto

© Copyright by Grace Beatrice Flock 2012

NOVEL RECEPTORS REGULATING INCRETIN SECRETION AND GLUCOSE HOMEOSTASIS

Grace Beatrice Flock

Doctor of Philosophy

Institute of Medical Science University of Toronto

2012

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Abstract

The ‐derived peptides (PGDP) are expressed in islet alpha and gut enteroendocrine L cells. Although , glucagon‐like peptide‐1 (GLP‐1), and glucagon like

peptide‐2 (GLP‐2) are derived from the same proglucagon , energy ingestion and nutrient assimilation represses proglucagon biosynthesis in the α‐cell, but stimulates the synthesis and

secretion of GLP‐1 and GLP‐2 from the gut L cell.

In the work presented in this thesis, I have identified novel G protein‐coupled receptors that stimulate GLP‐1 secretion and improve glucose homeostasis. G protein‐coupled

119 (GPR119) is expressed in enteroendocrine cells and islets and is activated by nutrients (fatty

acid derivatives) and small specific synthetic agonists. Activation of GPR119 enhances glucose‐ stimulated secretion from islet β‐cells and promotes incretin release from enteroendocrine cells in a cyclic AMP (cAMP)‐dependent manner. To determine the importance of gut for the glucoregulatory actions of GPR119, I examined GPR119 activation in normal mice, isolated islets, and in mice with inactivation of gut receptors. GPR119 activation directly stimulates insulin secretion from islets in vitro, yet requires intact incretin receptor signaling and enteral glucose exposure for optimal improvement of glucose tolerance in vivo. In contrast, activation of GPR119 inhibits gastric emptying independent of incretin receptors through GPR119‐dependent pathways.

Another important feature of β‐cell GPCRs coupled to cAMP generation is their ability to protect the β‐cell from external injury. I have shown that mice lacking GPR119 (GPR119‐/‐) are more susceptible to streptozotocin (STZ)‐induced apoptosis while pharmacological activation of

GPR119 failed to protect the β‐cell from STZ‐induced injury. Furthermore, GPR119‐/‐ mice iii

display impaired incretin secretion and function when chronically fed a high fat (HF) diet. Conversely, abrogation of GPR119 signaling does not affect the beta‐cell adaptation

(increased islet number and size) to the metabolic demand of high‐fat feeding.

Mechanisms to increase β‐cell mass and function may be useful tools for the treatment of

type 2 . GLP‐1 stimulates insulin biosynthesis, β‐cell proliferation and exerts anti‐ apoptotic actions on β‐cells. To delineate novel mechanisms, important for the regulation of proglucagon and GLP‐1 secretion in the enteroendocrine L‐cell, I carried out a microarray‐based gene expression profiling and transcriptional networks analysis using RNA

from murine gut GLUTag cells. To identify mechanisms unique to enteroendocrine L‐cells, I used the islet αTC1 cell line for comparative purposes. I identified a novel mediated signaling pathway involving activation of membrane GPCRs for the control of GLP‐1 secretion.

In summary, these studies establish that GPR119 engages multiple complementary pathways for control of glucose homeostasis and suggest that endogenous GPR119 signaling plays a critical role in β‐cell adaptation to cytotoxic injury and nutrient excess. The studies provide evidence for a novel role for progesterone, regulating GLP‐1 secretion and controlling

glucose homeostasis.

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Acknowledgments

First, I would like to start thanking my dearest grandfather, “abuelo Guillermo”, who has supported and encouraged me to follow my dreams. Without him, I would have not been able to start this amazing journey. Since the first days upon arriving to Canada, without speaking any English and with very limited resources, he has been my model, always in my mind and in my heart, reminding me that failure is never an option. I am also grateful to my parents who always believed in me and supported my endeavors.

I would like to give my sincere gratitude to my supervisor, Dr Daniel J. Drucker, who has supported and believed that it was important for me to complete my PhD studies. Without his support and understanding, becoming a Doctor in Philosophy would not have been possible. Furthermore, I am grateful to the members of my advisory committee, Dr Patricia Brubaker, Dr Adria Giacca and Dr Tony Lam for their thoughtful recommendations throughout my studies.

I also like to thank profoundly Dr Bernardo Yusta who had planted in my mind the seed to pursue and finish my PhD, a project that came to a stall many years earlier due to life circumstances. He has provided me with helpful discussions, and valuable advice. Bernardo had supported me with kind and encouraging words during troubled times as I was moving forward in this journey.

Many of the experiments could not have been done without the technical assistance of two very valuable members of Dr Drucker’s laboratory; Dianne Holland and Xiemin Cao and for that, I am very grateful to them.

I would like to thank former members of the lab; Ms Meghan Sauve, who had always been very kind and had given her time to assist me with several experiments, and Ben Lamont and Adriano Maida for their advice with a series of experimental protocols.

I am profoundly thankful to my dear friend, Theresa Kane, who did a masterful job in editing my thesis.

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Finally but not least, it is very important for me to extend my gratitude to my dear husband, Alberto, who has been a wonderful partner and friend supporting my studies, working overtime at home, looking after our beloved son, keeping the house together during those times when I was consumed by my work and responsibilities.

As well, I like to thank my son Axel, who has been so patient and understanding. Thanks Axel and remember that there is no limit for what you can become and never stop learning!

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Table of Contents

Abstract ...... iii

Acknowledgments ...... v

Table of Contents ...... vii

List of Tables ...... xi

List of Figures ...... xii

List of Abbreviations ...... xv

Author Contribution and Dissemination of Research ...... xx

Dedication ...... xxii

Chapter 1 INTRODUCTION ...... 1 1.1 GLUCAGON‐LIKE PEPTIDE‐1 ...... 2 1.1.1 The Incretin Effect ...... 2 1.1.2 The Proglucagon‐Derived Peptides...... 2 1.1.3 The GLP‐1 Receptor ...... 4 1.1.4 GLP‐1 actions, focus on the islet β‐cell ...... 7 1.1.5 GLP‐1 extra‐pancreatic effects ...... 13 1.2 GLUCAGON‐LIKE PEPTIDE‐1 (GLP‐1) SECRETION ...... 14 1.2.1 The Enteroendocrine System ...... 14 1.2.2 In vitro models for the study of GLP‐1 secretion regulation ...... 16 1.2.3 GLP‐1 secretion regulation ...... 19 1.2.4 Indirect regulation of GLP‐1 secretion: First phase ...... 20 1.2.5 Direct regulation of GLP‐1 secretion: Second phase ...... 26 1.3 GPR119 ...... 41 1.3.1 GPR119 expression ...... 41 1.3.2 GPR119 signal transduction ...... 41 1.3.3 GPR119 endogenous ligands ...... 42

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1.3.4 GPR119 insulinotropic actions ...... 43 1.3.5 GPR119 agonists and GLP‐1 secretion ...... 44 1.3.6 GPCRs and ‐cell cytoprotection ...... 45 1.3.7 GPR119 activation and regulation of gastric emptying and food intake ...... 46 1.4 NOVEL GPCRs MEDIATING GLP‐1 SECRETION ...... 48 1.4.1 Progesterone: genomic vs. non‐genomic actions ...... 48 1.4.2 Non‐receptor mediated non‐genomic progesterone actions ...... 49 1.4.3 Ion channel modulation mediating non‐genomic progesterone actions ...... 49 1.4.4 Nuclear receptors (PRs) mediating non‐genomic progesterone actions ...... 50 1.4.5 membrane component 1 and 2 (PGRMC1, PGRMC2) ...... 51 1.4.6 Novel GPCRs mediating non‐genomic progesterone actions ...... 51 1.5 RATIONALE FOR THE STUDIES DESCRIBED IN MY THESIS ...... 53 1.5.1 Chapter 2 ...... 53 1.5.2 Chapter 3 ...... 54 1.5.3 Chapter 4 ...... 55

Chapter 2 GPR119 REGULATES MURINE GLUCOSE HOMEOSTASIS THROUGH INCRETIN RECEPTOR‐DEPENDENT AND INDEPENDENT MECHANISMS ...... 57 2.1 ABSTRACT ...... 58 2.2 INTRODUCTION ...... 58 2.3 RESEARCH DESIGN AND METHODS ...... 60 2.3.1 Animal experiments ...... 60 2.3.2 Glucose tolerance tests ...... 60 2.3.3 Insulin Tolerance Test ...... 61 2.3.4 Arginine stimulation test ...... 61 2.3.5 Insulin secretion in mouse islets ...... 61 2.3.6 Analysis of GPR119 expression in islets ...... 62 2.3.7 Gastric emptying ...... 62 2.3.8 Statistical Analysis ...... 62 2.4 RESULTS ...... 63

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2.5 DISCUSSION...... 72

Chapter 3 GPR119‐/‐ MICE ARE MORE SUSCEPTIBLE TO STREPTOZOTOCIN‐INDUCED APOPTOSIS AND TO LIPOTOXIC‐INDUCED ‐CELL FAILURE ...... 79 3.1 ABSTRACT ...... 80 3.2 INTRODUCTION ...... 81 3.3 RESEARCH DESIGN AND METHOD ...... 82 3.3.1 Animal Experiments ...... 82 3.3.2 Streptozotocin‐Induced Apoptosis ...... 82 3.3.3 High Fat Diet Studies ...... 83 3.3.4 Glucose Tolerance Tests ...... 83 3.3.5 Insulin Tolerance Test ...... 84 3.3.6 Food Intake ...... 84 3.3.7 Magnetic Resonance Imaging (MRI) ...... 84 3.3.8 Indirect calorimetric and locomotor activity ...... 84 3.3.9 Islet Isolation ...... 85 3.3.10 Complementary DNA Synthesis and Gene Expression Analysis ...... 85 3.3.11 Pancreatic Insulin Content ...... 85 3.3.12 Islet Morphometric analysis ...... 86 3.3.13 Statistical analysis ...... 86 3.4 RESULTS ...... 86 3.5 DISCUSSION...... 104

Chapter 4 PROGESTERONE STIMULATES GLP‐1 SECRETION VIA MEMBRANE PROGESTERONE RECEPTORS YET IMPROVES GLUCOSE TOLERANCE VIA A GLP‐1R‐INDEPENDENT PATHWAY IN MICE ...... 110 4.1 ABSTRACT ...... 111 4.2 INTRODUCTION ...... 111 4.3 RESEARCH DESIGN AND METHODS ...... 113 4.4.1 Cell culture and microarray experiments ...... 113 4.4.2 RNA isolation and analysis ...... 114

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4.4.3 Cell transfection ...... 115 4.4.4 Signal transduction studies ...... 115 4.4.5 GLP‐1 levels ...... 116 4.4.6 Animal experiments ...... 116 4.4.7 Statistical analysis ...... 118 4.4 RESULTS ...... 118 4.5 DISCUSSION...... 132

Chapter 5 DISCUSSION...... 138

References ...... 148

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List of Tables

Chapter 3

Table 3.1 expressed at similar levels in GPR119‐/‐ compared to WT control islets…………91

Table 3.2 Islet mRNA transcripts with no change following HF feeding………………………………….103

Chapter 4

Table 4.1 Sequences for primers used to confirm microarray results…..……………...... 114

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List of Figures

Chapter 1: INTRODUCTION

Figure 1.1 Proposed mechanisms for how GPR119 activation regulates ‐cell function ...... 47

Chapter 2: GPR119 REGULATES MURINE GLUCOSE HOMEOSTASIS THROUGH INCRETIN RECEPTOR‐DEPENDENT AND INDEPENDENT MECHANISMS

Figure 2.1 GPR119 activation and control of oral glucose in WT and incretin receptor knockout mice...... 64

Figure 2.2 GPR119 activation and plasma levels of GIP, GLP‐1, insulin, and glucagon in WT mice during an intraperitoneal glucose tolerance test (IPGTT)...... 65

Figure 2.3 GPR119 activation and plasma levels of GIP, GLP‐1, insulin and glucagon in WT and incretin receptor knockout mice...... 66

Figure 2.4 GPR119 activation does not modify glucose profiles in an insulin tolerance test (ITT) in WT and DIRKO mice...... 68

Figure 2.5 GPR119 activation increases insulin secretion from WT and DIRKO islets...... 69

Figure 2.6 Gastric emptying in WT, Glp1r‐/‐, DIRKO, Gpr119‐/‐ and Glp2r ‐/‐ mice...... 71

Figure 2.7 GPR119 activation increases plasma PYY but inhibits gastric emptying independent of the Y2R receptor...... 73

Figure 2.8 The GPR119 agonist MBX3152 (Metabolex) failed to reduce food intake in mice ..... 77

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Chapter 3: GPR119‐/‐ MICE ARE MORE SUSCEPTIBLE TO STREPTOZOTOCIN‐INDUCED

APOPTOSIS AND LIPOTOXIC‐INDUCED ‐CELL FAILURE

Figure 3.1 Activation of GPR119 does not protect ‐cells from streptozotocin (STZ)‐induced apoptosis in WT mice; however, ablation of endogenous GPR119 signaling increases ‐cell susceptibility to STZ‐induced apoptosis...... 88

Figure 3.2 Levels of mRNA transcripts in islets isolated from GPR119‐/‐ and GPR119+/+ mice. 90

Figure 3.3 GPR119‐/‐ mice on a HFD develop fasting hyperglycemia and fail to control glucose homeostasis during OGTT...... 93

Figure 3.4 GPR119‐/‐ mice fed a HFD exhibit impaired ‐cell function...... 95

Figure 3.5 GPR119‐/‐ and WT control mice, chronically fed HFD do not display greater insulin resistance as measured by the insulin tolerance test (ITT)...... 96

Figure 3.6 No significant difference in islet size topography, number of islets, levels of insulin transcripts and pancreatic insulin content in GPR119‐/‐ compared to GPR119+/+ mice...... 97

Figure 3.7 No significant difference in the transcript levels of glucagon (Gcg), (PP) and (Sst) in of GPR119‐/‐ compared to GPR119+/+ following HF feeding...... 99

Figure 3.8 GPR119‐/‐ mice fed a HFD develop fasting hyperglycemia and fail to control glucose homeostasis during IPGTT...... 100

Figure 3.9 Islet levels of mRNA transcripts in GPR119‐/‐ and +/+ mice fed a HF diet as determined by qPCR...... 101

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Chapter 4: PROGESTERONE STIMULATES GLP‐1 SECRETION VIA MEMBRANE PROGESTERONE RECEPTORS YET IMPROVES TOLERANCE VIA A GLP‐1R‐INDEPENDENT PATHWAY IN MICE

Figure 4.1 Progesterone receptor is differentially expressed in GLUTag vs αTC1 cells. 120

Figure 4.2 The PR expressed in GLUTag cells is functional and transactivates the MMTV promoter; however, P4 does not modulate proglucagon gene transcription in vivo...... 121

Figure 4.3 Progesterone stimulates GLP‐1 secretion in a MAPK‐dependent manner in GLUTag cells...... 122

Figure 4.4 Progesterone stimulates GLP‐1 secretion via non‐genomic mechanisms activating Paqr5 and Paqr7 in GLUTag cells ...... 124

Figure 4.5 Progesterone stimulates GLP‐1 secretion in GLUTag cells independent of the classical progesterone receptor (PR)...... 126

Figure 4.6 Progesterone improves oral glucose tolerance and increases plasma GLP‐1 levels in mice...... 127

Figure 4.7 Progesterone lowers glucose and increases plasma GLP‐1 levels in mice via RU486‐ insensitive mechanisms...... 128

Figure 4.8 Progesterone improves oral glucose tolerance independent of incretin receptors. 129

Figure 4.9 Plasma incretin and insulin levels following enteral progesterone administration during OGTT...... 131

Figure 4.10 Acute enteral progesterone administration does not modulate the rate of gastric emptying and does not modify insulin sensitivity in mice...... 133

Figure 4.11 Intraperitoneally administered progesterone fails to improve glucose homeostasis during an OGTT in mice...... 134

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List of Abbreviations

18:1‐LPC Oleoyl‐lysophosphatidylcholine 2OG 2‐oleoyl glycerol 5‐HEPE 5‐hydroxy‐eicosapentaenoic acid AC Adenylyl cyclase ADP Adenosine‐5'‐diphosphate AKT1 serine/threonine‐protein kinase AR231453 Small synthetic molecule, specific GPR119 agonist (Arena Pharmaceuticals) AR881 Small synthetic molecule, specific GPR119 agonist (Arena Pharmaceuticals) ATG7 Autophagy‐related protein 7 ATP Adenosine‐5'‐triphosphate AUC Area under the curve BII0246 PYY receptor (Y2R) inhibitor Bip Heat shock 70kDa protein 5 (HSP5 also known as Bip) BSA‐P4 Bovine serum albumine‐progesterone conjugate 2+ Ca v Voltage dependent calcium channels CaGRP Calcitonin gene related peptide cal calories cAMP cyclic 3`, 5`‐adenosine monophosphate CBr receptor Cckr receptor cDNA complementary DNA CHO Chinese Hamster cell line CNS Central CO2 Carbon dioxide CREB Cyclic‐AMP response element binding protein DAG Diacylglycerol DIRKO Double incretin receptor knockout (homozygote) DMEM Dubelcco’s modified eagle medium DNA Deoxyribonucleic acid DPP‐4 Dipeptidyl peptidase‐4 EDTA Ethylenediaminetetraacetate EGFR Epidermal growth factor receptor ELISA ‐linked immunosorbent assay Epac Exchange protein directly activated by cAMP ER Endoplasmic reticulum ERK1/2 Extracellular signal‐regulated kinase Erα α FCS Fetal calf serum xv

Foxa2 Forkhead box protein A2 FoxO1 Forkhead 1 FRIC Fetal rat intestinal cells G protein Guanine nucleotide‐binding protein GABA Gamma Amino Butyric Acid GADD153 Growth Arrest and DNA Damage Inducible Protein 153 GDIR Glucose‐dependent insulinotropic receptor GDP Guanosine diphosphate GIP Glucose‐dependent insulinotropic peptide GIPR Glucose‐dependent insulinotropic peptide receptor Gipr‐/‐ Glucose‐dependent insulinotropic peptide receptor knockout mice GIRK G‐protein‐regulated inward rectifying K+ channels GK Glucokinase GLP‐1 Glucagon‐like peptide‐1 Glp1r Glucagon‐like peptide‐1 receptor (gene) GLP‐1R Glucagon‐like peptide‐1 receptor (protein) Glp1r‐/‐ Glucagon‐like peptide‐1 receptor knockout mice Glp2 Glucagon‐like peptide‐2 Glp2r‐/‐ Glucagon‐like peptide‐2 receptor knockout mice GLUT1 Facilitative glucose transporter‐5 GLUT2 Glucose transporter‐2 GLUT5 Fructose transporter GLUTag Murine enteroendocrine L‐cell line GPCR G protein‐coupled receptor GPR119 G protein‐coupled receptor 119 GPR119‐/‐ G proetin‐coupled receptor 119 knockout mice GRP ‐releasing peptide Grpr Gastric releasing peptide receptor (RNA) GSIS Glucose‐stimulated insulin secretion GTP Guanosine triphosphate HCl Hydrochloric acid HEK293 Human embryonic kidney cell line HF High fat HFD High fat rodent diet (45% Kcal from fat) HPMC Hypromellose HRE Hormone response elements HSP Heat shock protein Iapp IGF‐1 Insulin‐like growth facor IGF‐1R Insulin‐like growth facor receptor Igf‐2r Insulin‐like growth facor receptor‐2 (gene) IL‐6 Interleukin‐6 IP Inositol phosphate

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ip Intraperitoneal IP‐2 Intervening peptide‐2 IPGTT Intraperitoneal glucose tolerance test IRS2 Insulin receptor substrate 2 IRS‐2 Insulin receptor substrate‐2 Kcnj11 potassium inwardly‐rectifying channel subfamily J, member 11 Kv Delayed rectifier channel LPE Lysophosphatidylethanolamine LPI MAPK Mitogen‐activated protein kinase MBX3152 Small synthetic molecule, specific GPR119 agonist (Metabolex) MH Meat hydrolase MIN6 Murine insulinoma cell line mPR Membrane‐bound progesterone receptor mRNA messenger RNA MUFA Monounsaturated fatty acids NCI‐H716 Human enteroendocrine L‐cell line NeuroD Neurogenic differentiation factor NIT‐1 Murine β‐cell line N‐terminal amino‐terminal OEA OGTT Oral glucose tolerance test OLDA Oleoyldopamine P4 Progesterone P62 Sequestosome 1 (gene = SQSTM1, protein = P62) PACAP Pituitary adenylate cyclase‐activating polypeptide PAQR Progestins and adiponectin receptor family PC Prohormone convertase PC1/3 Pro‐hormone convertases 1/3 PC2 Pro‐hormone convertase 2 PCR Polymerase chain reaction PD98059 Inhibitor of active MEK1,2 PDX‐1 Pancreas 1 PGDPs Proglucagon derived peptides PGRMC1/2 Progesterone receptor membrane component 1 and 2 PI3K Phosphatidylinositol‐3 kinase PKA Protein kinase A PKB Protein kinase B (also known as Akt) PKC Protein kinase C PKCζ Atypical protein kinase C‐zeta PLC Phospholipase C PP Pancreatic polypeptide PPAR Peroxisome proliferator‐activated receptor

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PR Progesterone receptor (classical) PRA isoform PRB isoform PSN632408 Small synthetic molecule, specific GPR119 agonist (Prosedium ) PYY Peptide YY qPCR Quantitative polymerase chain reaction RC Standard rodent chow diet (18% Kcal from fat) RH7777 Rat hepatoma cell line RIA Radioimmunoassay RIMS2 Regulating synaptic membrane exocytosis protein 2 RNA Ribonucleic acid RT Reverse transcriptase RT‐PCR reverse transcriptase‐PCR RU486 Progesterone antagonist RyR Ryanodine receptors SGLT‐1 Sodium‐dependent glucose transporter‐1 SGLT‐3 Sodium glucose transporter‐3 siRNA small interfering ribonucleic acid SS Somatostatin SS‐14 Somatostatin‐14 SS‐28 Somatostatin‐28 STC‐1 Murine enteroendocrine L‐cell line STZ Streptozotocin SUR1 Sulfonylurea Receptor Type 1 Subunits TC Tissue culture TCF7L2 Transcription factor 7‐like 2 TRVP1 Vanilloid‐responsive transient receptor potential vanilloid 1 UCP2 Mitochondrial uncoupling protein 2 UO126 Inhibitor of both active and inactive MEK1, UPR Unfolded protein response V Vehicle Vapb vesicle‐associated membrane protein‐associated protein B vol volume Vpac‐1 Vasoactive Intestinal Peptide Receptor‐1 Vpac‐2 Vasoactive Intestinal Peptide Receptor‐2 vs versus WT Wild‐type Y2R PYY receptor‐2 Munc‐18 Mammalian uncoordinated‐18 EMSA Electrophoretic mobility shift assay ChIP Chromatin Immunoprecipitation

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Methodological abbreviations % percent oC degrees Celsius Da Dalton g gram h hour(s) l litres M molar (moles/l) min minute(s) mol moles sec second(s) U units vol volume wt Weight

Prefixes k kilo‐ (x 10‐3) c centi‐ (x 10‐2) m milli‐ (x 10‐3) μ micro‐ (x 10‐6) n nano‐ (x 10‐9) p pico‐ (x 10‐12)

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Author Contribution and Dissemination of Research

Chapter 1: INTRODUCTION Contribution: Grace B. Flock produced all text for this chapter. Dr D.J. Drucker advised with editing the text for this chapter.

Publication: No part of the text in this chapter has been reproduced elsewhere.

Chapter 2: GPR119 REGULATES MURINE GLUCOSE HOMEOSTASIS THROUGH INCRETIN RECEPTOR‐DEPENDENT AND INDEPENDENT MECHANISMS Contribution: Dianne Holland contributed to animal experiments, providing technical assistance (Figures: 2.1 and 2.3). Grace B. Flock generated data for all figures in this chapter, performed data analysis, and figure preparation. Dr D.J. Drucker advised with experimental design. G.B. Flock and D.J. Drucker wrote the paper together.

Publication citation: (2011). 152(2):374‐83.

Chapter 3: GPR119 NULL MICE ARE MORE SUSCEPTIBLE TO STREPTOZOTOCIN ‐INDUCED APOPTOSIS AND TO LIPOTOXIC‐INDUCED ‐CELL FAILURE Contribution: Grace B. Flock generated data for all figures in this chapter, performed data analysis, and figure preparation. Dr D.J. Drucker advised with experimental design and figure preparation. G.B. Flock produced all text in this chapter. Dr D.J. Drucker advised with editing the text for this chapter.

Publication: No part of the text in this chapter has been reproduced elsewhere.

Chapter 4: PROGESTERONE STIMULATES GLP‐1 SECRETION VIA MEMBRANE PROGESTERONE RECEPTORS YET IMPROVES GLUCOSE TOLERANCE VIA A GLP‐1R‐INDEPENDENT PATHWAY IN MICE Contribution: Xiemin Cao contributed to experiments designed to confirm microarray results (Figure 3.1A) and to the measurement of glucagon levels during IPGTT (Figure 4.10C). Marlena Maziarz contributed with bioinformatics based microarray analysis. Grace B. Flock generated

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data for all remaining figures in this chapter, performed data analysis, and figure preparation. Dr D.J. Drucker advised with experimental design and figure preparation. G.B. Flock and Dr D.J. Drucker wrote this chapter together.

Publication: No part of the text in this chapter has been reproduced elsewhere.

Chapter 5: DISCUSSION Contribution: Grace B. Flock produced all text for this chapter. Dr D.J. Drucker advised with editing the text for this chapter.

Publication: No part of the text in this chapter has been reproduced elsewhere.

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Dedication

To the memory of my grandfather

Wilhelm J. Flock

En memoria de mi abuelo

Guillermo J. Flock

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1

Chapter 1

INTRODUCTION

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1.1 GLUCAGON‐LIKE PEPTIDE‐1 1.1.1 The Incretin Effect The incidence of type 2 diabetes mellitus, a metabolic disorder characterized by high blood glucose in the context of insulin resistance, is rising worldwide [1]. It is estimated that 330 million people will have diabetes by the year 2030 [2]. Over 100 years ago, it was hypothesized that there were factors released by the intestine that were able to promote the secretion of internal components of the pancreas thereby reducing the circulating levels of glucose [3, 4]. By the middle of the 1960s, it was shown that glucose administered orally promoted a much greater insulin release from the pancreas than when administered intravenously [5, 6]. It is now widely recognized that the oral ingestion of glucose triggers a phenomenon known as the “incretin effect” which accounts for 50% to 70% of the total insulin release. The incretin effect is largely attributed to two hormones secreted by enteroendocrine cells, glucagon‐like peptide‐1 (GLP‐1) and glucose‐dependent insulinotropic peptide (GIP) [7‐11]. Under normal physiological conditions is believed that both GIP and GLP‐1 contribute equally to the incretin effect. However, diabetic patients exhibit a reduced incretin effect. This deficiency is believed to be the result of reduced GLP‐1 secretion and impaired GIP action [12‐14].

1.1.2 The Proglucagon‐Derived Peptides The mammalian gene encoding proglucagon is transcribed as a single messenger RNA (mRNA), expressed in three distinct cell types: the intestinal enteroendocrine L‐cells, the α‐cells of the endocrine pancreas and neurons located in the and brainstem [15, 16]. The transcription of the proglucagon gene in α‐cells and L‐cells is differentially regulated. In the islet α‐cell, fasting and hypoglycemia up‐regulates and insulin inhibits proglucagon gene expression. Conversely, fasting reduces and feeding enhances proglucagon mRNA levels in the intestine [17]. Transcription factors such as Brn4, Pax6 and members of the forkhead transcription factor family are involved in the control of proglucagon gene transcription[18‐20]. The Wnt signaling pathway via binding of the transcription factor TCF‐4 (currently named TCF7L2) to the proglucagon gene promoter has been shown to up‐regulate proglucagon gene transcription in rodent enteroendocrine L‐cells but not in islet α‐cells [21]. Translation of the pre‐proglucagon

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mRNA transcript results in a 160 amino acid precursor. Posttranslational processing by endo‐ proteases, named pro‐hormone convertases, gives rise to proglucagon‐derived peptides (PGDPs) [22]. Tissue‐specific expression of different pro‐hormone convertases dictates the resulting PGDPs that are expressed in that tissue. Hence, in the α‐cells of the endocrine pancreas pro‐hormone convertase 2 (PC2) is partially responsible for the cleavage of proglucagon to generate, predominantly, glucagon as well as glicentin‐related polypeptide (GRPP), intervening peptide‐1 (IP‐1), and the major proglucagon fragment (MPGF). In the enteroendocrine L‐cell pro‐hormone convertases 1/3 (PC1/3) will generate glicentin, , intervening peptide‐2 (IP‐2), glucagon‐like peptide‐1 and glucagon‐like peptide‐ 2 [23‐25]. In contrast, it is still not well established which pro‐hormone convertase is responsible for the processing of proglucagon in the central nervous system. High levels of both PC1/3 and PC2 have been localized in the hypothalamus, where neurons expressing proglucagon can also be found [26]. Several studies have demonstrated the production of small amounts of GLP‐1 in the pancreatic ‐cell [27‐29]. For example, a significant increase of PC1/3 expression leading to an increase in GLP‐1 synthesis and secretion from islet ‐cells was demonstrated in human islets maintained in the presence of high glucose concentrations for a prolonged period of time [30]. Similar results were seen in rodents where diabetes was induced [30, 31] and in models of murine ‐cell regeneration [32]. Studies using isolated rat islets have shown that the processing of proglucagon in ‐cells switches to increase GLP‐1 production as a response to ‐cell glucotoxicity or chemical‐induced injury [33]. Active GLP‐1 exists in two forms, GLP‐1 (7‐37) and GLP‐1 (7‐36) amide. GLP‐1 (7‐36) amide represents the major active circulating form in humans [34], while GLP‐1 (7‐37) predominates in pigs, dogs and rats [35]. Both forms of GLP‐1 contain an alanine in position 2, which renders them substrates for the proteolytic action of dipeptidyl peptidase‐4 (DPP‐4). Hence, GLP‐1 (7‐ 37) and GLP‐1 (7‐36) amide are rapidly degraded to GLP‐1 (9‐37) and GLP‐1 (9‐36) amide having a half‐life of 2 min [36, 37]. Moreover, GLP‐1 has been shown to be a substrate for neutral endopeptidase 24.11 (NEP 24.11) in vitro [38]. NEP 24.11 is a membrane‐bound Zn metallopeptidase that cleaves peptides at the N‐terminal side of aromatic or hydrophobic

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amino acids. Six potential cleavage sites have been identified in the C‐terminal of the GLP‐1 molecule. The truncated peptides were thought to be inactive; however there is a growing body of evidence supporting the involvement of GLP‐1 (9‐36) amide in glucose clearance and cardiovascular functions [39‐43]. Two types of incretin‐based therapies have been developed for the treatment of diabetes: GLP‐1 receptor agonists and DPP‐4 inhibitors which prolonged GLP‐1 and GIP activity via inhibition of DPP‐4 [44]. Exendin‐4, a peptide isolated from the of the Heloderma suspectum lizard [45] is a potent GLP‐1R agonist [46], exhibits 53% sequence identity with the human GLP‐1 and its actions in rodents and humans resemble those of GLP‐1 (reviewed in [47]). Exendin‐4 is resistant to the proteolytic actions of DPP‐4, thus it possess a bioactive half‐life of 3 to 4 hours in humans [48]. In 2005, the FDA approved exendin‐4 (Byetta®) for the treatment of type 2 diabetes. Moreover, new strategies are currently in the pipeline involving the development of incretin secretagogues, which in combination with DPP‐4 inhibitors could result in therapeutic tools of greater potency for the treatment of diabetes.

1.1.3 The GLP‐1 Receptor The GLP‐1R first cloned from human and rat islet cDNA libraries [46], is a class B G‐protein coupled receptor (GPCR) [49]. The rat and human GLP‐1R share 90% amino acid identity [46]. Consistent with the general structure of a GPCR, the GLP‐1R is composed of seven transmembrane domains linked by 3 intracellular and 3 extracellular loops, an intracellular C‐ terminal region and an extracellular amino‐terminal region [50]. GPCRs, upon binding, undergo conformational changes that will allow the receptor to

bind G‐proteins and transduce the signal. G‐proteins are composed of three subunits; Gα, G β and

Gγ. In the inactive state, Gα bounds to guanosine diphosphate (GDP) and is associated to the G βγ

heterodimer. Following receptor activation, GDP‐bound Gα is exchanged for guanosine triphosphate (GTP)‐bound Gα resulting in the release of the G βγ complex. Downstream signaling

is initiated by Gα and the G βγ complex [50]. There are four subfamilies of Gα proteins: Gαs, Gαi

which includes Gαo, Gαq and Gα12.

The effector of both the Gαs and Gαi/o pathways is the cyclic‐adenosine monophosphate (cAMP) generating enzyme adenylyl cyclase (AC). AC catalyzes the conversion of cytosolic

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adenosine triphosphate (ATP) to cAMP. While Gαs activates AC resulting in an increase in cAMP

formation, Gαi/o inhibits AC activity and cAMP formation [51, 52]. The effector of the Gαq/11 pathway is phospholipase C (PLC), which catalyzes the cleavage of membrane‐bound phosphatidylinositol 4, 5‐diphosphate (PIP2) into the second messengers inositol (1, 4, 5) triphosphate (IP3) and diacylglycerol (DAG). IP3 acts on IP3 receptors found in the membrane of the endoplasmic reticulum (ER) to elicit Ca2+ release from the ER, while DAG diffuses along the plasma membrane where it may activate any membrane‐localized forms of a ser/thr kinase called protein kinase C (PKC) [52]. Since many isoforms of PKC are also activated by increases in intracellular Ca2+, both these pathways can also converge on each other to signal through the same secondary effector [53]. The Gα12 family plays a role in the regulation of cytoskeletal assembly [51].

The primary effectors of Gβγ are various ion channels, such as G‐protein‐regulated inward rectifying K+ channels (GIRKs), P/Q and N‐type voltage‐dependent Ca2+ channels, as well as some isoforms of AC and PLC, along with some phosphoinositide‐3‐kinase (PI3K) isoforms [53].

GLP‐1R has been shown to couple to Gs, Gαq, Gαi and Gαo. The best‐characterized actions of

the pancreatic GLP‐1R arise from its coupling to Gs, activation of AC and the subsequent increase of cAMP production [8, 54]. Downstream signaling involves activation of protein kinase A (PKA) and of exchange protein directly activated by cAMP (Epac) [8, 55]. Treatment of murine islets with the PKA inhibitor H89 or down‐regulation of Epac2 expression levels with antisense oligonucleotide reduced, but did not prevent, GLP‐1‐mediated insulin secretion. In contrast, co‐ administration of H89 and Epac oligonucleotide antisense completely abrogated the secretagogue action of GLP‐1 in mouse islets [56]. Thus, GLP‐1 utilizes both PKA and Epac signaling pathways to stimulate insulin secretion. For example, studies in murine βTC6 and hamster HIT‐T15 insulinoma cell lines and in rat islets demonstrated that GLP‐1, via PKA and Epac activation, induces elevation of intracellular calcium levels [57] and potentiates glucose‐ stimulated insulin secretion (GSIS). The increase in intracellular calcium involves activation of voltage‐dependent calcium channels localized to the plasma membrane of β‐cells, as well as inducing the release of calcium from intracellular stores [57‐59]. Heterologous expression of the rat GLP‐1R in a monkey kidney cell line (COS‐7) demonstrated that GLP‐1R also couples to Gαq, leading to PLC activation and increased

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intracellular IP3. This induces mobilization of intracellular calcium and activation of classical and novel PKC isoforms [60]. Studies in insulinoma cell lines and transfected heterologous cells have shown that GLP‐1R

can also couple to Gαi and Gαo [61, 62]. Moreover, studies in INS‐1 cells demonstrated that the proliferative actions of GLP‐1 are dependent on betacellulin‐mediated activation of the epidermal growth factor receptor (EGFR) [63]. GLP‐1 transactivates EGFR through the activation of c‐Src and the subsequent activation of a membrane‐bound metalloproteinase and the concomitant release of betacellulin [63‐65]. EGFR transactivation by GLP‐1 is followed by activation and translocation to the nucleus of the protein kinase C atypical isoform zeta (PKCζ), resulting in enhanced β‐cell proliferation [65]. GLP‐1 has also been shown to activate ERK1/2 in a glucose‐dependent manner, in Chinese hamster ovary (CHO) cells stably expressing rat GLP‐1R, isolated rodent islets and Ins1 and Min6 cell lines [61, 66]. This effect was inhibited by pretreatment with cholera toxin (CTX) in

CHO cells expressing the rat GLP‐1R [61, 66]. It was also mimicked by forskolin and blocked by H89 in MIN6 cells. Thus, GLP‐1 mediated ERK1/2 activation is cAMP‐PKA dependent [67]. In addition, ERK1/2 activation by GLP‐1 was shown to require both an increased influx of calcium via L‐type voltage‐gated calcium channels and the activation of calcium/calmodulin‐dependent protein kinase II [67]. The proliferative and antiapoptotic actions of GLP‐1 on islet β‐cells have been shown to be partially dependent on GLP‐1‐mediated activation of the phosphatidylinositol 3‐kinase (PI3K)‐AKT signal transduction pathway [68‐71]. In humans and rodents, the GLP‐1R has been shown to be expressed in insulin‐producing ‐ cells, glucagon‐producing ‐cells, somatostatin‐producing ‐cells and in the pancreatic ducts [72‐74]. GLP‐1R expression was also shown in lung, heart, intestine, kidney, and , neurons of the nodose ganglion of the vagus, skin, and several regions of the CNS. Furthermore, while GLP‐1R expression has been demonstrated in canine muscle and adipose tissue, its expression in rodent and human muscle, liver and adipose tissue remains a subject of debate [75‐77].

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1.1.4 GLP‐1 Actions, Focus on the Islet β‐cell Glucose sensing and insulin secretion It is believed that glucose enters the ‐cell via the high capacity, low affinity glucose transporter 2 (GLUT2) to be rapidly phosphorylated by glucokinase (GK). The low affinity of GK enables regulation of glycolytic flux over a physiological range of glucose concentrations [78]. Hence, GK is the rate‐limiting factor for glycolysis in ‐cells. As ambient glucose concentrations rise, for example after a meal, intracellular glucose concentrations will increase to a threshold that will favor GK‐mediated glucose phosphorylation. Phosphorylated glucose will then enter the glycolytic pathway. Glucose oxidation generates an increase in Adenosine‐5'‐triphosphate (ATP) production with a concomitant change in the ATP/ADP ratio. The increase in ATP/ADP + ratio will trigger the closure of ATP sensitive K channels (KATP) resulting in membrane depolarization. The change in opens the voltage‐dependent calcium 2+ 2+ channels (Ca v) with the consequent influx of Ca . Increased concentrations of intracellular calcium will then stimulate insulin exocytosis [79‐81], reviewed in [82].

GLP‐1 actions on glucose sensing and insulin secretion GLP‐1 was first identified as an insulinotropic peptide able to stimulate insulin secretion in a glucose‐dependent manner [59, 83]. As described under the GLP‐1R signal transduction section, cAMP and the downstream activation of PKA and Epac2 are the main pathways involved in the insulin‐secreting actions of GLP‐1 [8, 55]. Furthermore, prolonged GLP‐1R activation restores glucose sensitivity in previously resistant β‐cells [84]. This is an effect that could be partially explained by the ability of GLP‐1R signaling to induce the expression of glucose transporters and glucokinase, which as discussed above are critical components of the β‐cell glucose sensing machinery [85]. GLP‐1 in addition to increasing the levels of insulin secreted per β‐cell [86] sensitizes the β‐ cell to ambient glucose levels via its ability to modulate the activity of the ATP‐dependent K+

channel (KATP) and the delayed rectifier channel (Kv); channels critical to insulin secretion [87,

88]. KATP channels consist of four Kir6.2 units that form the pore of the channel and four SUR1 regulatory subunits. ATP binds to the Kir6.2 subunit causing a conformational change that induces inhibition of the channel activity [89]. Studies in the Kir6.2 knockout mouse [90] and

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the SUR1 knockout mouse [91] demonstrated that closure of KATP channels and membrane depolarization are critical for first phase insulin secretion. In humans, mutations in Kir6.2 or SUR1 subunits result in severe hypoglycemia associated with hyperinsulinemia [92]. GLP‐1 increases the glucose‐dependent membrane depolarization by facilitating the closure of KATP channels [84, 93]. It is believed that PKA‐dependent and Epac2‐dependent phosphorylation of

SUR1 subunits facilitates the closure of KATP channels in rodent and human β‐cells [93, 94].

Voltage‐dependent channels (Kv) are responsible for membrane repolarization and the

termination of action potentials. Many Kv subfamilies have been identified in mammals [95, 96]. In the resting β‐cell, Kv channels are closed. Upon membrane depolarization and following glucose‐induced insulin secretion, Kv channels open [87]. The opening of Kv channels generates a voltage‐dependent K+ outward current that restores resting membrane potential. The importance of modulating the activity of Kv channels, specially the Kv2.1 member, for glucose‐ stimulated insulin secretion was demonstrated in studies in rat islets where 60‐70 % reduction of Kv2.1 activity by expressing a dominant negative form resulted in a 60% increase in insulin secretion [97]. Patch‐clamp experiments in rat islets demonstrated that GLP‐1 and exendin‐4 antagonize Kv currents [98]. Thus, GLP‐1R activation prolongs islet β‐cell electrical activity by

inhibiting the voltage‐dependent rectifying channels (Kv) [99]. This action is mediated via cAMP/PKA pathway and via transactivation of the EGF receptor and the downstream PI3K/PKCζ pathways [98, 100]. Increased intracellular calcium is a critical event for the exocytosis of insulin vesicles, reviewed in [101‐103]. Intracellular calcium rises via two main mechanisms triggered by activation of AC and GLP‐1 signaling. One mechanism involve the activation of voltage‐ dependent calcium channels following membrane depolarization, causing them to open and allowing an influx of extracellular Ca2+. A second mechanism involves the enhancement of calcium‐induced Ca2+ release from intracellular stores. GLP‐1R signaling, via PKA activation, induces voltage dependent L‐type Ca2+ channels phosphorylation, increasing their open capability and enhancing Ca2+ influx [104]. GLP‐1 treatment also induces release of Ca2+ from the endoplasmic reticulum through two intracellular‐Ca2+ releasing receptors; the inositol 1,4,5

triphosphate receptor (IP3R) activated by cAMP‐ PKA and the ryanodine receptors (RyR) activated by the cAMP‐Epac2 dependent pathway [105, 106].

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Oxidation of pyruvate, a product of glucose metabolism, is critical for insulin secretion. Pyruvate enters the β‐cell mitochondria to be metabolized via the KREB cycle. Intra‐ mitochondrial Ca2+ concentrations regulate the activity of several dehydrogenases involved in this metabolic process, including pyruvate dehydrogenase. Studies in the mouse insulinoma cell line MIN6, using bioluminescence imaging of adenoviruses expressing mitochondrial‐targeted luciferase facilitated measurement of cytosolic versus mitochondrial ATP. Similarly, using adenoviruses expressing the Ca2+‐dependent photoprotein, mitochondrial aequorin, allowed measuring mitochondrial Ca2+ levels [107]. In these studies, the GLP‐1‐induced increase in calcium‐induced calcium release from intracellular stores resulted in an increase in mitochondrial Ca2+ and ATP levels above the levels induced by glucose alone [106].

GLP‐1 effects on insulin exocytosis In endocrine cells, regulated exocytosis of large dense‐core vesicles is a process that involves vesicle recruitment to the plasma membrane, docking of vesicles at the plasma membrane, priming of fusion machinery and fusion of the vesicles with the plasma membrane. An increased intracellular calcium level is an essential event for regulated exocytosis. In addition, diacylglycerol (DAG), ATP and phospholipids also modulate Ca2+‐induced exocytosis [101, 108]. cAMP enhances translocation of insulin granules to the plasma membrane, increases the size of the readily releasable pool and promotes the rate of replenishment of insulin granules via cAMP‐Epac dependent mechanisms [102, 109]. In addition to its effects on membrane depolarization, GLP‐1 is believed to regulate exocytosis by enhancing these steps via PKA‐dependent and PKA‐independent mechanisms [59, 99, 110]. PKA phosphorylates proteins associated with the exocytotic machinery of β‐cells including mammalian uncoordinated‐18 (Munc‐18) and soluble N‐ethylmaleimide‐sensitive factor attachment protein (α‐SNAP) [111]. In addition, GLP‐1 activation of AC induces the dissociation of cAMP‐ Epac2 from the Sur1 subunits 2+ of the KATP channel [94]. Free cAMP‐Epac2 promotes Ca ‐ dependent dimerization of the vesicle‐associated proteins Rim2 and Piccolo. Rim2‐piccolo complex interacts with a core component of the exocytotic system, the binding protein Rab3A [101].

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GLP‐1 effects on insulin biosynthesis In addition to potentiation of insulin secretion, GLP‐1 also promotes, together with glucose, insulin mRNA and protein synthesis [8, 112, 113] in a PKA‐dependent and ‐independent manner [8, 114]. Studies in cultures of rat insulinoma RIN 1046‐38 cell line treated with GLP‐1 demonstrated an increase in insulin mRNA transcript levels [8]. Studies in rat INS‐1 cells stably transfected with expression reporter vectors, containing the human insulin gene promoter provided further evidence for GLP‐1‐mediated activation of the human insulin promoter [115]. The effect of GLP‐1R activation on insulin protein synthesis was illustrated treating isolated rat islets with exendin‐4. Using metabolic pulse‐radiolabel techniques followed by immunoprecipitation and PAGE, it was demonstrated that GLP‐1R activation potentiates glucose‐stimulated insulin protein biosynthesis [113].

GLP‐1 effects on β‐cell mass Beta cell mass is the result of a balance between β‐cell proliferation and β‐cell death. GLP‐1 is a potent β‐cell growth factor that exhibits proliferative, neogenic and anti‐apoptotic action. The proliferative properties of GLP‐1 were demonstrated in pancreatic mouse and rat cell lines and in cultures of rodent islets [70, 116, 117]. Furthermore, continuous administration of GLP‐1 to Zucker diabetic fatty rats (ZDF), which exhibit a defect in the leptin receptor and therefore overeat, become obese, insulin resistant and diabetic, demonstrated increased β‐cell proliferation compared to vehicle‐treated animals [118]. Studies in mice exhibiting a defect in the leptin receptor (db/db mice) also demonstrated that GLP‐1R activation leads to increased β‐ cell proliferation in young rodents [119]. GLP‐1 was shown to increase DNA synthesis as measured by the incorporation of tritiated thymidine in insulinoma cell lines and in rat [70]. This action of GLP‐1 was shown to be mediated via transactivation of the epidermal growth factor receptor (EGFR) followed by PI3K‐AKT mediated phosphorylation and nuclear translocation of the atypical isoform protein kinase C zeta (PKCζ) [63]. Exactly how PKCζ nuclear translocation leads to increased β‐cell proliferation is not completely understood. Another signaling pathway involved in GLP‐1 mediated induction of β‐cell proliferation is the activation of AKT (also named PKB). GLP‐1 signaling activates AKT via the EGFR‐PI3K and the cAMP‐PKA‐CREB‐IRS2 signaling pathways, reviewed in [120]. Forkhead transcription factor 1 (FoxO1) is a transcription factor implicated mediating β‐cell growth induced by insulin and

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insulin‐like growth factor‐1 (IGF‐1) signaling [121]. The activity of FoxO1 is inhibited by AKT‐ dependent phosphorylation and nuclear exclusion in INS‐1 cells. Inactivation of FoxO1 induces pancreas duodenum homeobox 1 (Pdx‐1) nuclear translocation and the transcriptional up‐ regulation of Pdx‐1 synthesis. Pdx‐1 is a critical transcription factor for pancreatic development [122], and is essential for GLP‐1‐mediated β‐cell trophic effects [123]. Cell cycle proteins tightly control G1/S transition in cell cycle progression [124]. Cell cycle progression is critical for the post‐natal maintenance of β‐cell mass. β‐cells express cyclin D1 and cyclin D2. Absence of cyclin D2 in mice results in the development of diabetes in cyclin D2‐/‐ mice [125]. GLP‐1 and exendin‐4 treatment have been shown to induce the mRNA expression of cyclin D1 via activation of PKA, PI3K and MEK/ERK signaling pathways in rodent insulinoma cell lines [117, 126]. In fact, the cyclin D1 promoter contains a CRE element and using electrophoretic mobility shift assay (EMSA) and chromatin Immunoprecipitation (ChIP) analysis in INS‐1 cells it was demonstrated that exendin‐4 increases the association of phospho‐ CREB to the CRE element of the cyclin D1 promoter [126] . Additionally, GLP‐1 proliferative actions involve activation of the Wnt signaling pathway in β‐cells [127]. In INS‐1 cells, both GLP‐ 1 and exendin‐4 stimulated luciferase activity driven by transcription factor 7‐like 2 (TCF7L2) consensus‐binding sites. Furthermore, exendin‐4 induced PKA‐dependent phosphorylation of β‐ catenin allowing it to interact with TCF7L2 and activate Wnt genes such as cyclin D1 [127]. Interestingly, mRNA levels of TCF7L2 were upregulated, and protein levels were significantly reduced in isolated islets from rodent models of type 2 diabetes compared with the non‐ diabetic controls. Furthermore, the mRNA expression level of GLP‐1R was decreased in islets from humans with T2DM as well as in isolated human islets treated with siRNA to TCF7L2 [128]. Moreover, common variants of TCF7L2 are associated with the development of type 2 diabetes [129, 130]. GLP‐1 also augments IGF‐1R expression and induces IGF‐2 secretion enhancing the IGF1‐R/IGF2 autocrine loop that leads to GLP‐1 induced β‐cell proliferation [131]. GLP‐1 has been shown to prevent β‐cell death both in vitro [69, 71, 132] and in rodent models of β‐cell injury [118, 119, 132]. Studies using the low dose streptozotocin (STZ)‐induced β‐cell destruction model [132] and cytokine‐induced apoptosis [131] in mice lacking the GLP‐1R provided evidence for a role of GLP‐1 signaling protecting islet β‐cells from injury. In these studies, Glp1r‐/‐ mice were shown to be more susceptible to STZ‐induced and cytokine‐induced

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β‐cell death. Furthermore, exendin‐4 protected WT mice from STZ‐induced β‐cell death [132]. Perfetti et al demonstrated that GLP‐1 protected MIN6 cells and cultured human islets from reactive oxygen species (ROS)‐induced apoptosis [71, 133]. Using pharmacological inhibitors, the PI3K and cAMP pathways were shown to be critical in GLP‐1 cytoprotective actions from ROS‐induced injury, whereas the MAPK pathway played no role [71]. GLP‐1 was also shown to exhibit cytoprotective actions against β‐cell lipotoxicity both in vitro [134] and in vivo [119]. Western blot analysis of pancreatic homogenates from db/db mice treated with exendin‐4 exhibit an increased expression of AKT1, an isoform expressed selectively in β‐cells, and ERK1, compared to vehicle‐ treated animals [119 4196]. AKT (PKB) activation has been shown to protect β‐cells from lipotoxicity [135]. In accordance, the cytoprotective actions of GLP‐1 from free fatty acid‐ induced apoptosis in β‐cells is mediated via AKT (PKB) activation and possibly via its downstream target nuclear factor ΚB [134]. Similar to the proliferative effects of GLP‐1, the anti‐apoptotic actions are also mediated via EGFR and PI3K‐ dependent activation of AKT (PKB) followed by phosphorylation and nuclear exclusion of FoxO1 and the consequent nuclear translocation and up‐regulation of PDX‐1 expression. Moreover, cAMP‐PKA‐dependent activation of CREB also mediates the anti‐apoptotic actions of GLP‐1 by up‐regulation of IRS2 expression, and the consequent activation of AKT, reviewed in [120, 136]. In addition, GLP‐1 via enhancing the IGF1‐R/IGF2 autocrine loop and the downstream phosphorylation of AKT exhibited protective actions from cytokine‐induced β‐cell death in the MIN6 cell line and mouse islets [131].

GLP‐1 effects on the secretion of other islet hormones GLP‐1 also exerts actions in islet α‐cells inhibiting glucagon secretion in a glucose‐ dependent manner. Furthermore, GLP‐1 stimulates somatostatin release from islet δ‐cells even under normoglycemic conditions [137, 138] . It has been proposed that the glucagonostatic action of GLP‐1 could be mediated by direct activation of a putative GLP‐1R on α‐cells [72], or indirectly via the paracrine actions of insulin and/or somatostatin release [139]. GLP‐1‐ mediated somatostatin release is believed to be mediated via direct activation of the GLP‐1R expressed on δ‐cells [140].

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1.1.5 GLP‐1 Extra‐Pancreatic Effects For the purpose of this introduction, I will briefly discuss GLP‐1 actions in the intestinal tract and the nervous system as they relate to the studies presented in this thesis.

Gastrointestinal tract: GLP‐1 effect on gastric emptying GLP‐1 not only stimulates glucose‐dependent insulin secretion, but also improves postprandial glucose control via complementary mechanisms. GLP‐1 inhibits the rate of gastric emptying via inhibition of gut motility, thus slowing the transit of nutrients from the to the intestine and reducing postprandial glucose excursions even in diabetic patients [141, 142]. GLP‐1 actions on gut motility are mediated primarily via neural pathways particularly the vagus nerve as demonstrated by studies in rats that underwent vagal afferent denervation and in humans that had truncal vagotomy, where the inhibitory actions of GLP‐1 on gut motility were completely abrogated [143, 144].

Nervous system: GLP‐1 effect on food intake and satiety Neurons expressing GLP‐1 as well as neurons expressing the GLP‐1R are found in regions of the brain that control appetite, behavior, gastric motility, glucoregulation and cardiovascular functions. Intracerebroventricular and peripheral administration of GLP‐1 and exendin‐4 have been shown to reduce food intake and promote weight loss in normal and obese rodents [145, 146]. Intra‐venous or subcutaneous administration of GLP‐1 in humans produced a dose dependent reduction in food intake [147‐149]. Furthermore, GLP‐1 and exendin‐4 promoted satiety and reduction of body weights in obese and diabetic individuals [150‐152]. The hypothalamic arcuate nucleus may be involved in GLP‐1 regulation of food intake as chemical destruction of this region abolishes GLP‐1‐induced anorectic actions [153]. In addition, a study in rats that were cannulated at the arcuate nucleus and injected with GLP‐1 or saline suggested that GLP‐1 signaling via the arcuate nucleus is not involved in its ability to modulate food intake [154]. Similar to what has been seen for GLP‐1 actions on gastric emptying, ablation of the afferent vagal circuits abolished the anorectic actions of peripherally administered GLP‐1 and exendin‐4 in rats [155]. Moreover, peripherally administered GLP‐1 failed to induce c‐Fos‐like immunoreactivity in the arcuate nucleus of vagotomised rats [155], while administration of an albumin‐bound GLP‐1 protein, a large molecule unable to cross the

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blood‐brain barrier, activated neurons in the rodent brain [156], thus suggesting the involvement of vagal afferent signaling to transduce GLP‐1 actions in the arcuate nucleus. The GLP‐1R is also expressed on cells of the nodose ganglion of the afferent vagus [157]. Very little active GLP‐1 reaches the pancreas due to rapid inactivation by DPP‐4 [158]. Thus, it is believed that GLP‐1 may exert its insulinotropic actions via activation of neuronal pathways in vivo. In fact, ganglionic blockade abolished the increase in glucose‐stimulated insulin release by intraportal GLP‐1 administration in rats [159].

1.2 GLUCAGON‐LIKE PEPTIDE‐1 (GLP‐1) SECRETION 1.2.1 The Enteroendocrine System GLP‐1 is released by L‐cells, specialized open‐type enteroendocrine cells that make direct contact with the intestinal lumen and sense and respond to nutrients [160]. The enteroendocrine system is the largest endocrine organ in the body. The intestine is essentially a tube organized in four concentric layers: the mucosa, the , the muscularis external and the serosa. The mucosa is the inner layer surrounding the intestinal lumen and it refers to the combination of the epithelium plus the . The lamina propria is a thin layer of loose connective tissue that lies beneath the epithelium. The mucosa is responsible for the processing and absorption of nutrients [161], for secretory processes as well as for the compaction of the stools. The submucosa is composed by a dense irregular layer of connective tissue containing large blood vessels, lymphatic and nerves that branch into the mucosa and the external muscularis external. The muscularis external is composed of several sheets of smooth muscle and enteric nerve fibers that control the peristaltic movements of the intestine [162]. The outermost layer is the serosa and consists of several layers of connective tissue. The intestine can be divided anatomically in two main sections, the , and the , which can be subdivided from proximal to distal in the duodenum, and . The absorptive surface of the small intestine is greatly increased by the formation of epithelial protrusions into the lumen, the villi, and embedding into the sub‐mucosa, the crypts of Lieberkuhn. In contrast, in accordance with its major role in stool compaction rather than nutrient absorption the large intestine’s mucosa consists of an inner simple columnar

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epithelium that lacks villi and presents deeper embedding of the crypts into the sub‐mucosa. Its main functions are to complete the final absorption (less than 5%) of nutrients from food, to reabsorb the water added to the chyme by the secretions of the digestive system, and to compress and compact the remaining wastes into feces. The maintenance and repair of the epithelial lining of the intestine is achieved by a mechanism of self‐renewal where proliferation is balanced with apoptosis and shedding [163, 164]. The epithelial lining of the human intestine renews every 5‐7 days. This process is dependent on the self‐renewal and pluripotent characteristics of the adult stem cell that resides deep within the intestinal crypt; reviewed in [165]. These stem cells give rise to all cell types found in the intestinal mucosa [163]. The intestinal mucosa contains four distinct cell types that differ in the relative distribution and abundance throughout the intestine, reviewed in [166, 167]: are the most abundant cells in the epithelial layer and are responsible for the absorption of nutrients and the synthesis of the glycoprotein needed for terminal and absorption. They include disaccharidases, and enteropeptidase, which converts (inactive) trypsinogen to (active) trypsin. Trypsin, in turn, activates trypsinogen itself, as well as other pancreatic zymogens that facilitate nutrient absorption [166, 168]. Goblet cells are simple columnar epithelial cells that secrete mucin, which dissolves in water to form . They increase in abundance from the small to the large intestine. Secreted mucus protects the intestine from shear stress and chemical damage. Paneth cells control the intestinal by secreting antimicrobial agents such as cryptidins (defensins in humans) and lysozyme [169] and are largely restricted to the crypts of the small intestine. Enteroendocrine cells are found scattered among the enterocytes in the epithelial layer throughout the intestinal tract [166]. They represent a small portion of the entire intestinal epithelia (less than 1%). They are either closed or open types of cells depending on whether they are buried in the mucosa (closed type) or open to the lumen (open type). Open type cells are conically shaped, with protrusions (microvilli) emerging from the apical pole and extending into the intestinal lumen and with a larger base where peptide hormones will be released from dense secretory granules. Many different sub‐types of enteroendocrine cells have been

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described based on immunological and morphological differences and the corresponding list of gut hormones that they produce is even larger [170]. Among these cells, we find the GLP‐1‐ secreting L‐cell and the GIP‐secreting K‐cell. It is noteworthy to mention the existence of less well defined cell types including brush/tuft cells, cup cells, and the M cells localized on lymphoid Peyer’s patches in the intestine [166]: Brush/Tuft cells possess varied morphological appearance. Their biological functions have not been completely defined. These cells are found in the ducts of the salivary glands and have been shown to express α‐gustducin and transient receptor potential cation channel subfamily M member 5 (TRPM5), critical components of the signaling system, indicating a possible role as chemosensors. However, on the basis of morphological, histochemical and cytochemical evidence, secretory and absorptive functions have also been proposed. For example, tuft cells may underlie the release of opioids into the gut lumen following nutrient ingestion, which in turn exhibit effects on gastric emptying modulation and intestinal secretions [171], for a review see [172]. Cup‐cells are specialized non‐absorptive cells of unknown function found in the [173]. M‐cells are specialized epithelial cells. They are polarized with a typical apical and basolateral membrane. The basolateral membrane however presents an intraepithelial pocket that acts as the docking site for intraepithelial lymphocytes [174‐176]. M‐cells are associated with Peyer's patches and lymphoid follicles. They actively take up macromolecules and microorganisms from the intestinal lumen and are thought to be the portal of entry for bacteria and viruses [177].

1.2.2 In Vitro Models for the Study of GLP‐1 Secretion Regulation The study of GLP‐1 secretion in native mature L‐cells has been difficult due to the low abundance of L cells. Thus, our current knowledge derives from experiments in which hormone secretion was measured in the whole body or using vascular or luminal perfusion models [178]. The study of molecular mechanisms and signaling pathways involved in the regulation of GLP‐1 secretion relied on in vitro systems like primary cultured fetal enteroendocrine cells [179] and immortalized endocrine cell lines like human NCI‐H716, and murine GLUTag and STC‐1 [180‐

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184]. Next, I will provide a brief description of these valuable tools for the study of GLP‐1 secretion and the mechanisms involved.

NCI‐H716 This cell line was derived from a spontaneous human cecal adenocarcinoma and secretes GLP‐1 [184]. However, even though these cells are able to respond to a limited number of secretagogues, the control of proglucagon gene expression and gene transcription differs from data obtained in studies using primary rodent intestinal cell cultures. For example, activation of the adenylate cyclase pathway, known to increase proglucagon gene expression in islets and enteroendocrine rat and murine cell models, did not increase the levels of proglucagon‐mRNA transcripts in human NCI‐H716 cells [185]. Moreover, NCI‐H716, transfected with reporter plasmids containing from 328 pair of bases (pb) to over 5700 pb of human proglucagon promoter and 5’ flanking sequence, were transcriptionally inactive. Conversely, the same reporter plasmids were shown to be transcriptionally active when transfected in islet InR1‐G9 cells [185]. RT‐PCR analysis demonstrated that NCI‐H716 cells express several proglucagon gene transcription factors important for rodent proglucagon gene expression, including Pax6 and ‐2. In contrast, mRNA transcripts for Brn‐4, a positive regulator of proglucagon gene transcription in rodent islets, was not detected in the same study. Despite this limitation, they are extensively used as a model for studies of human enteroendocrine PGDP secretion.

Fetal Rat Intestinal Cells The establishment of primary cell cultures utilizing cells enzymatically dispersed from 19‐ to 21‐day fetal rat intestines (FRIC) was instrumental to help delineate the mechanisms that control GLP‐1 secretion [179].These cells were shown to secrete GLP‐1 and respond to a variety of secretagogues [179, 180, 182]. However, FRIC cultures are a heterogeneous population of primary cells that do not secrete GLP‐1 in response to glucose, a well‐known stimulus that promotes GLP‐1 secretion. An explanation could be that these “fetal” cells have not yet differentiated to fully express the glucose sensing machinery [179, 186].

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GLUTag cells This is a homogeneous stable immortalized cell line, derived from a colonic glucagon‐ producing enteroendocrine cell tumor that arose from glucagon gene‐SV40 T antigen transgenic mice [187]. They express the proglucagon gene and retain the ability to secrete GLP‐1 in response to a variety of secretagogues; hence, they are extensively used to gain an understanding of L‐cell biology [183, 186‐194]. Moreover, GLUTag cells retain many of the properties associated with differentiated gut endocrine cells, including cAMP‐dependent regulation of proglucagon gene transcription and the ability to sense and respond to changes in nutrient availability. Primary fetal rat cell cultures (FRIC) were used to confirm the validity of GLUTag cells as a model for the study of GLP‐1 secretion. [188].

STC‐1 This cell line derived from a mouse duodenal tumor resulting from a cross between rat insulin promoter‐ SV40 large T antigen (RIP‐Tag) and rat insulin promoter polyoma small T antigen (RIP‐PyST) transgenic mice [195]. These cells are plurihormonal, expressing genes for the duodenal hormones and cholecystokinin (CCK), as well as the proglucagon gene [196, 197]. They have been used to study GLP‐1 biosynthesis. However, these cells display an aberrant posttranslational processing of proglucagon resulting in significant quantities of glucagon being produced in addition to glicentin, oxyntomodulin and GLP‐1 [198‐200]. Nevertheless, the STC‐1 cells have been used in studies of GLP‐1 and GIP biosynthesis and secretion, and have been shown to be responsive to both protein hydrolysate mix and cholinergic agonists [201‐203].

Native L‐cells isolated from the adult mouse Recent advances have allowed the identification, isolation and study of native L cells from the adult mouse [193]. Bacterial Artificial (BAC) constructs were generated to express a modified yellow fluorescent protein (YFP‐Venus) under the control of the proglucagon gene promoter (Venus was cloned in place of the coding region of proglucagon, between the start codon in exon 2 and the stop codon in exon 6). These constructs were used to generate transgenic mice. Primary cultures of small intestine and colon were then generated from these

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mice. Venus positive cells were easily identified providing the opportunity to study murine mature native L‐cell biology, electrophysiology and signal transduction pathways [193]. Fluorescence Activated Cell Sorting (FACS) followed by microarray analysis allowed for the transcriptome profiling of adult native L cells. Comparison of the transcriptome of mature native L‐cells and GLUTag cells demonstrated that they exhibit very similar patterns of gene expression for key receptors, glucose sensors, ion channels, and regulatory molecules known to be important in GLP‐1 secretion regulation [193]. Primary cultures of native L‐cells from adult mice have confirmed the validity of GLUTag cells as a model for the study of the enteroendocrine L‐cell biology [193, 194, 204, 205]. Interestingly, microarray analysis has also revealed differences between colonic and duodenal murine adult L cells. While colonic L‐cells highly express GLP‐1, PYY and GLP‐2, they express very low levels of CCK and they do not co‐express GIP. However, L cells from the duodenum express high levels of CCK, and a very small sub‐population co‐express very high levels of GIP [193]. This is contrary to the classical thought that GIP and CCK were produced and secreted from distinct enteroendocrine cells other than the L‐cell. These new discoveries indicate a more complex and plastic enteroendocrine system than was previously thought. GLUTag cells, in accordance with the site of generation of this cell line, were shown to resemble the colonic murine L‐cell [193].

1.2.3 GLP‐1 Secretion Regulation The L‐cell, at the apical membrane, presents protrusions (microvilli) that extend into the lumen of the intestinal tube. The microvilli allow the L‐cell to sense luminal signals and respond accordingly [160, 193]. Due to its localization, the basolateral membrane of L‐cells is in close proximity to nerves and vascular tissue [160]. As a result, GLP‐1 secretion from the L‐cells is regulated by multiple neural, hormonal and nutrient signals. The main physiological stimulus for GLP‐1 secretion is the ingestion of nutrients. Carbohydrates and fat are the most potent stimuli, while proteins appear to be less potent [206, 207]. Studies have shown that there is a “caloric threshold” needed to trigger GLP‐1 secretion from the L‐cell [208, 209]. Peak values of nutrient‐stimulated GLP‐1 are dependent on the rate at which those nutrients are delivered into the duodenum [210] and the load of nutrients [211].

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Levels of GLP‐1 are low in the fasting state [207] and, in humans, rise by two to three fold after a mixed meal [206, 208, 212]. Furthermore, studies in healthy individuals have helped to delineate the pattern of GLP‐1 release. GLP‐1 is released tonically, in a pulsatile manner with a frequency of 5 to 7 pulses per hour, under basal conditions [213] . Oral glucose administration increases circulating GLP‐1 levels; a consequence of an increase in GLP‐1 pulse amplitude but not of pulse frequency of secretion [213]. In vivo, nutrient‐stimulated GLP‐1 secretion occurs in a biphasic manner, consisting of a rapid early phase within 5‐15 minutes after food ingestion and lasting for 15‐30 min [206, 214, 215], followed by a more sustained second phase approximately 60‐90 minutes later [216]. Enteroendocrine L‐cells can be found throughout the small and the large intestine [160, 193, 217] with greater density in the ileum and colon [160, 218]. Hence, the rapid rise in the levels of GLP‐1 after a meal occurs before nutrients have reached the majority of the GLP‐1 producing enteroendocrine cells [160, 219‐221]. A few enteroendocrine cells that co‐express GLP‐1 and GIP have been found in the duodenum [193, 222]. These duodenal L‐cells could account for the rapid rise in GLP‐1 induced by nutrients. However, studies in humans [213] and rats [223‐225] support the concept of an indirect nutrient‐mediated GLP‐1 secretion stimulation via a duodenal‐ileal‐neuro‐endocrine loop. For example, in rats, preventing the transit of nutrients to the ileum by duodenal ligation demonstrated that fat load into the duodenum stimulated GLP‐1 secretion. The increase of GLP‐1 was the result of activation of the ileal L‐cells since removal of the distal gut abrogated the nutrient‐induced GLP‐1 secretion [225, 226].

1.2.4 Indirect Regulation of GLP‐1 Secretion: First Phase Neuronal Control A strong body of evidence has established an important role for the vagus nerve, the adrenosympathetic system, and the neurotransmitters acetylcholine, calcitonin gene‐related peptide (CGRP), gastrin‐releasing peptide (GRP) and Gamma Amino Butyric Acid (GABA), as mediators of the proximal‐distal control of GLP‐1 secretion from the ileal L‐cell. For example, direct electrical stimulation of the celiac branch of the vagus nerve that innervates the jejunum, ileum and colon, resulted in stimulation of GLP‐1 secretion in rats [223]. Left cervical vagotomy of anesthetized rats resulted in a significant reduction of basal circulating

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GLP‐1 levels demonstrating a role for the vagus nerve in regulating tonic release of GLP‐1 under non‐stimulated conditions [223]. Conversely, bilateral sub‐diaphragmatic vagotomy in the rat demonstrated that no other nerve circuits other than the vagus mediate the proximal‐distal loop. In this model, corn oil infusion into the isolated duodenum failed to promote GLP‐1 secretion [223]. The vagus nerve releases acetylcholine, which is an agonist for both nicotinic and muscarinic receptors. Studies in humans [213] and in rats [224] demonstrated that atropine, a non‐specific muscarinic receptor antagonist, diminishes the nutrient‐induced early rise of GLP‐1 indicating an essential role for cholinergic‐muscarinic signaling in the control of the L‐cell. Using fetal rat intestinal cell cultures (FRIC), murine GLUTag and human NCIH716 cell lines it was demonstrated that activation of muscarinic receptors M1 and M2 results in stimulation of GLP‐ 1 secretion [188, 224, 227, 228]. Furthermore, double immunostaining of the rat ileum with antibodies against GLP‐1, and the muscarinic receptor isoforms M1, M2 and M3 showed co‐ expression of GLP‐1 and all three subtypes of muscarinic receptors [224]. However, using specific antagonists to all three isoforms, it was demonstrated that corn oil‐induced GLP‐1 secretion is dependent on M1 receptor activation in rats [224]. Studies in rats also demonstrated a role for nicotinic receptors mediating the early rise of GLP‐1 release. For example, blockade of nicotinic receptors by infusions of hexamethonium, a nicotinic receptor antagonist that has no effect on muscarinic receptors, prevented the effect of intraduodenal oleic acid administration in mediating GLP‐1 secretion [229].

The adrenosympathetic system The adrenosympathetic system and its effectors have been shown to induce hormone secretion from enteroendocrine L cells [216, 230, 231]. Adrenosympathetic neurotransmitters act through three receptors, 1 and 2. Stimulation of adrenergic receptors by infusions with epinephrine in the isolated perfused rat ileum model induced significant GLP‐1 and PYY secretion which was abolished by the non‐selective beta blocker propranolol and strongly decreased upon infusion with the alpha adrenergic blocker prazosin [232]. In contrast, blockade of alpha2‐receptors with idazoxan did not alter epinephrine‐induced peptide secretion. Thus, GLP‐1 secretion is positively modulated by stimulation of adrenergic receptors β and α1 while is inhibited by activation of α2 receptors [232].

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Calcitonin gene‐related peptide (CGRP) Local afferent fibers are extensions of vagal afferent fibers and directly innervate the local target cell without synapsis to central centers. They contain the neurotransmitter CGRP that has been shown to stimulate GLP‐1 secretion in vitro [227, 231, 233]. It has been proposed that local sensory afferent nerves in the may play a role in the timing of early GLP‐1 release [223, 233].

Gastric Releasing Peptide (GRP) The duodenum is innervated by vagal afferents that are responsive to nutrients, particularly fat [234]. Immunoreactive GRP localizes in gastrin‐releasing peptide‐containing nerve cell bodies in the myenteric ganglia all along the gut [235, 236]. Mice lacking a functional GRP receptor exhibit reduced glucose‐induced circulating levels of GLP‐1, decreased insulin release and they are glucose intolerant when subjected to an oral glucose challenge [237]. Studies in anesthetized rats demonstrated a role for GRP mediating fat‐ induced GLP‐1 secretion. In these studies, infusion of the GRP antagonist BW10 completely prevented the GLP‐1 response to duodenal fat from the distal L‐cell [225]. Studies using the isolated perfused rat ileum model also demonstrated a role for GRP in the regulation of PGDPs [216, 231]. Furthermore, studies in enteroendocrine human (NCIH716) cells and rodent FRIC cultures and GLUTag cells have demonstrated that GRP is a potent GLP‐1 secretagogue [216, 225, 227, 231].

Gamma Amino Butyric Acid (GABA) GABA is a neurotransmitter that stimulates GLP‐1 secretion in vitro. GABA receptor is a chloride channel that, when activated, promotes the efflux of negatively charged chloride ions. Studies in GLUTag cells demonstrated that GABA triggers membrane depolarization and GLP‐1 release [192]. However this finding was in contrast with work performed by other groups where GABA was not found to be a secretagogue in both rat perfused intestine and GLUTag cell cultures [188, 216]. Nevertheless, whether GABA stimulates GLP‐1 secretion from the native colonic murine L‐cell has not been investigated.

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Endocrine Control In addition to neuronal control, a series of hormones play an important role in the indirect mechanisms that regulate nutrient‐stimulated GLP‐1 secretion.

Glucose‐dependent insulinotropic peptide (GIP): GIP is released from enteroendocrine K‐ cells following ingestion of fat and/or carbohydrates [238, 239]. The highest density of K‐cells is localized to the duodenum [240, 241]. In rodents, physiological levels of GIP stimulate GLP‐1 secretion from the distal L‐cell via the engagement of vagal circuits in a GRP‐dependent manner [242]. Using fetal rat intestinal cells (FRIC) [179] and murine GLUTag cell cultures [188, 226, 227] it was demonstrated that supraphysiological concentrations of GIP directly stimulates GLP‐ 1 secretion. Studies using the isolated vascularly perfused rat ileum preparation, where different peptides and neurotransmitters were infused, demonstrated that GIP mediates GLP‐1 secretion from the rodent ileum [216, 231]. Interestingly, infusion of GIP into rats administered as a bolus through the jugular vein followed by continued infusion through the femoral vein, stimulated GLP‐1 secretion independent of glucose levels [242]. In contrast, in hepatic branch vagotomized rats, infusion of GIP no longer was able to stimulate GLP‐1 from the distal L‐cell [223]. Conversely, when GIP was infused at supraphysiological concentrations, GIP stimulated GLP‐1 secretion independent of the presence of an intact vagal circuit [223]. Hence, at least in rodents, physiological levels of GIP mediate GLP‐1 secretion from the distal L‐cell via the engagement of vagal circuits, while

supraphysiological levels are able to directly stimulate the L‐cell. GIP‐mediated modulation of GLP‐1 secretion is species‐specific. In pigs, GIP can stimulate GLP‐1 secretion only when administered in supraphysiological concentrations but fails to do so in physiological postprandial concentrations [243]. In humans, infusions of GIP failed to increase GLP‐1 secretion [13] and no correlation between post prandial levels of GIP and GLP‐1 have been found [209]. Histological data from human intestine suggests the presence of similar number of L‐cells and K‐cells in the duodenum [244]. Furthermore, there are enteroendocrine cells that co‐ secrete GLP‐1 and GIP in the proximal portion of the small intestine [193, 244], thus opening the possibility of a model for a GIP‐mediated paracrine regulation of GLP‐1 secretion. However,

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the observation that in humans GLP‐1 levels increased prior to increases in GIP after oral nutrient ingestion [207, 209] argue against a paracrine regulation, thus further studies will be necessary to support this concept. Cholecystokinin (CCK) is a gastrointestinal peptide released from intestinal I cells in response to long chain fatty acids and proteins via activation of GPR40 and extracellular calcium‐sensing receptors respectively [245‐247]. CCK actions include inhibition of gastric emptying and food intake, stimulation of pancreatic enzyme release together with increasing gallbladder contractions, reviewed in [248]. A study in healthy male volunteers that were administered an intraduodenal infusion of sodium oleate together with IV infusion of saline or the CckAr antagonist DEXLOX demonstrated a role for CCK release and CckAr activation in the fat‐mediated modulation of GLP‐1 secretion[249]. Intraduodenal infusions of sodium oleate together with IV saline significantly increased circulating levels of CCK and GLP‐1 release. In contrast, GLP‐1 secretion was attenuated when sodium oleate was co‐administered with DEXLOX [249]. Thus, CCK release, via activation of CckAr, appears to play an important role in the modulation of fat‐induced GLP‐1 secretion in humans. Insulin: In humans, a population‐based study has found that insulin resistance is associated with impaired GIP and GLP‐1 responses to nutrients [250]. Furthermore, an association between impaired glucose tolerance, impaired insulin response to oral glucose and reduced

GLP‐1 release has also been demonstrated [251]. The MKR mice that express a dominant negative IGF‐1R in the muscle and develop insulin resistance in muscle, adipose tissue and liver [252] exhibit impaired GLP‐1 secretion when administered an oral glucose load [253]. The insulin receptor is expressed in human and murine L‐cells [253]. In vitro, insulin activates PI3K‐ AKT and Ras‐Raf‐MAPKK‐p44/42 MAPK pathways and promotes GLP‐1 secretion from FRIC, GLUTag and NCI‐H716 enteroendocrine L‐cell models [253]. However, prolonged exposure of enteroendocrine cell cultures to insulin resulted in impaired GLP‐1 release, [253]. Hence, it has been postulated that the L‐cell is responsive to insulin and could become insulin resistant during the development of glucose intolerance and progression to type 2 diabetes [253]. In GLUTag cells, insulin stimulates actin remodeling via activation of the Rho guanosine 5’‐ triphosphatase cell division cycle 42 (Cdc42) and the downstream Cdc42‐dependent p21‐ activated kinase‐1 (PAK1), an event required for insulin‐mediated GLP‐1 secretion [254].

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Furthermore, knockdown or inhibition of both Cdc42 and PAK1 prevented ERK1/2 phosphorylation by insulin, which as indicated above, is a critical pathway for insulin‐mediated GLP‐1 release [253, 254]. Leptin is a cytokine released from the adipose tissue and functions as the afferent signal acting on receptors in the hypothalamus of the brain where it inhibits appetite [255]. In addition, it is also locally produced in the stomach [256]. For example, immunohistochemistry studies detected leptin immunoreactive cells in normal human gastric samples [256]. RIA studies detected leptin in human gastric juices [256]. Furthermore, both leptin and leptin receptor mRNA transcripts were detected by RT‐PCR in the human stomach [256]. In fasted rats, gastric leptin is released 15 min after re‐feeding [257]. Intraperitoneal administration of recombinant leptin to rats and ob/ob mice, resulted in a significant increase in the levels of circulating GLP‐1 [258, 259]. In vitro, leptin was shown to increase GLP‐1 secretion directly from FRIC, GLUTag and NCI‐H716 cells [259]. is associated with hyperleptinemia. It has been postulated that the impaired GLP‐1 secretion observed in human obesity [260] could be a consequence of the development of L‐ cell leptin resistance. Evidence to support this hypothesis comes from studies where C57Bl‐6 mice were fed a high fat (45%) or low fat (10% of Kcal from fat) diet for eight weeks. The mice fed the high‐fat diet (HFD) became obese, developed insulin resistance, hyperinsulinemia, hyperglycemia and hyperleptinemia [259]. Moreover, these mice developed leptin resistance as assessed by measuring food consumption following recombinant leptin administration. In addition, mice on HFD had lower fasting and glucose‐stimulated plasma GLP‐1 levels compared to lean mice [259]. Interleukin‐6 (IL‐6) is a cytokine that can exert pro‐inflammatory and anti‐inflammatory actions. Il‐6 is produced by immune cells, such as monocytes and lymphocytes, in adipose tissue and in the muscle [261, 262]. In adipose tissue, most of IL‐6 is not released by mature adipocytes but rather by preadipocytes, endothelial cells and monocytes‐macrophages [263]. IL‐6 production by adipose tissue increases in obesity [264, 265]. In obese subjects, elevated IL‐ 6 is believed to be associated with the development of insulin resistance and diabetes [264‐ 266]. In contrast, exercise has been shown to acutely promote IL‐6 release from skeletal muscle, improving peripheral insulin sensitivity [267‐269].

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IL‐6 was shown to promote islet α‐cell proliferation and prevent α‐cell apoptosis [270]. Furthermore, using IL‐6 KO mice it was demonstrated that IL‐6 signaling is critical for HFD‐ mediated expansion of α‐cell mass in vivo [270]. Interestingly, IL‐6 KO mice fed a HFD also exhibited impaired postprandial insulin secretion [270]. Acute intraperitoneal (ip) administration of IL‐6 and exercise‐mediated increase in IL‐6 levels, both improved oral glucose excursions and insulin secretion in a GLP‐1‐dependent manner in mice [271]. Furthermore, elevation of IL‐6 increased pancreatic GLP‐1 content in mice and stimulates GLP‐1 synthesis and secretion from GLUTag cells [271]. Somatostatin (SS) is produced as two isoforms: somatostatin‐14 (SS‐14) and somatostatin‐ 28 (SS‐28). While SS‐14 is mainly of neuronal origin, SS‐28 has been found to be produced by enteroendocrine D‐cells [272, 273]. Studies using the isolated perfused porcine ileum preparation demonstrated that GLP‐1 secretion is tonically inhibited by a local release of SS‐28 [274]. Studies using FRIC cultures provided direct evidence for somatostatin‐mediated inhibitory actions of GLP‐1 secretion, [227]. Since GLP‐1 stimulates somatostatin (SS) release from the enteric D‐cell [275], it has been postulated that both hormones operate under an auto‐regulatory system.

1.2.5 Direct Regulation of GLP‐1 Secretion: Second Phase The direct interaction of luminal nutrients with enteroendocrine L‐cells is most likely of fundamental importance during the second phase of GLP‐1 secretion. Enteroendocrine cells, like islet β‐cells are equipped with nutrient‐sensory machinery. Nutrient‐sensing will activate a variety of intracellular signaling pathways including membrane depolarization, elevation of intracellular Ca2+ levels and second messengers like cAMP, PLC, MAPK cascades.

G‐protein coupled receptors (GPCRs) regulating GLP‐1 secretion

Gs‐Coupled receptors cAMP is a potent stimulus for GLP‐1 secretion in NCI‐H716 [184], GLUTag [276], FRIC [188, 277] and canine enteroendocrine [230] cell cultures. In GLUTag cells, cAMP acts synergistically with glucose and directly stimulates GLP‐1 secretion via a mechanism that involves membrane depolarization and stimulation of the secretory machinery [276]. cAMP levels are regulated by a balance between adenylate cyclase (AC)‐mediated production and phosphodiesterase‐

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mediated hydrolysis. In primary colonic L‐cells the combination of the general AC activator forskolin with the non‐selective phosphodiesterase inhibitor 3‐isobuty‐1‐methylxanthine (IBMX) induced GLP‐1 secretion synergistically with glucose and associated with an increase in intracellular calcium levels [193, 278].

CGRP and GIP have been shown to stimulate GLP‐1 secretion via Gαs activation [188, 242,

243]. The fatty acid receptor GPR119 and the acid receptor TGR5 are GPCRs coupled to Gs and they have been shown to modulate GLP‐1 secretion. These receptors will be discussed in more detail under the Fatty Acid Receptor section on page 36.

Gq‐mediated GLP‐1 secretion

Muscarinic agonists bind to Gq‐coupled receptors and stimulate incretin secretion in rats and in FRIC, GLUTag and NCI‐H716 cell cultures [224, 228]. Bombesin and gastrin‐releasing peptide (GRP) also signal via Gq‐coupled receptors and stimulate GLP‐1 secretion [180, 202,

279]. The fatty acid receptors GPR40, GPR120 and GPR43 couple to Gq and are postulated to modulate GLP‐1 secretion. They will be discussed in more detail under the Fatty Acid Receptors section on page 36.

Gi‐coupled receptors

Somatostatin has been shown to inhibit GLP‐1 secretion though activation of the Gi‐coupled subtype 5 (SSTR5) in FRIC cultures [277]. The fatty acid receptor GPR41 couples to Gi and it is discussed in more detail later on under the Fatty Acid Receptors section on page 36.

Carbohydrate‐mediated GLP‐1 secretion As previously discussed, glucose sensing has been extensively studied in pancreatic ‐cells where insulin is released in response to an increase of ambient glucose levels; reviewed in [82]. Remarkably, in vitro studies have demonstrated that glucose‐mediated secretion pathways in L‐ cells and ‐cells, for the most part, share common glucose‐sensing machinery [186]. Studies in GLUTag cells [186] and in primary intestinal cultures of adult murine L‐cells [204] demonstrated that the L‐cell is an electrically active entity able to sense and respond to glucose. For example, intact GLUTag cells and primary murine L‐cells were studied using the perforated‐patch clamp

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method. It was demonstrated that at low concentrations of glucose (0 – 2 mM) these cells are for the most part quiescent and hyperpolarized. Upon glucose stimulation, they undergo electrophysiological changes resulting in membrane depolarization triggering action potentials and GLP‐1 secretion [186, 193]. GLUTag cells express glucokinase (GK) transcripts, however cytoplasmic extracts obtained from them did not display glucokinase activity over a range of glucose concentration [191]. In fact, most of the detected activity was due to hexokinase [191, 193]. GLUTag cells were shown to express the high‐affinity hexokinases I‐III [191]. Interestingly, humans with mutations in the glucokinase gene develop onset of diabetes early on in life (MODY‐2) however, they do not display abnormal incretin secretion [280].

RT‐PCR and microarray studies demonstrated the presence of K‐ATP channel subunits Kir6.2 and SUR1 in human NCI‐H716 and in murine GLUTag and native L‐cells [190, 193]. Immunostaining detected Kir6.2 in L and K cells of the human intestine [244, 281]. Tolbutamine, an agent that closes K‐ATP channel, triggered action potentials associated with increased in intracellular calcium and stimulated GLP‐1 secretion in GLUTag and primary L‐cell cultures. Conversely, diazoxide, an agent that opens ATP sensitive K+ channel, had the opposite effect

[186, 193] thus, confirming a functional role for K‐ATP channels for the transduction of glucose‐ mediated GLP‐1 secretion in L‐cells. Despite the presence of K‐ATP channels in humans and mouse L‐cells, sulfonylurea (agents that bind to the sulfonylureas receptors (SUR‐1) subunit of the K‐ATP channel promoting its closure) have failed to significantly increase levels of GLP‐1 in humans [282]. Moreover, patients with K‐ATP channel mutations that cause the channel to remain opened at basal ATP concentrations display early onset of diabetes due to defective insulin secretion, but they show no alteration in GLP‐1 levels[283]. These findings suggests that despite the presence, for the most part, of the molecular glucose sensing machinery used by ‐ cells, in GLUTag cells this does not seem to be the dominant mechanism that couples carbohydrate sensing to GLP‐1 secretion. As with islet β‐cells, glucose‐mediated GLP‐1 secretion correlates with increase in intracellular calcium levels in GLUTag and native L‐cells. Microarray studies demonstrated the presence of the L‐type and N‐type voltage dependent Ca2+ channels in GLUTag cells and L‐type Ca2+ channels in native L‐cells. Patch clamp studies and secretion studies have shown that

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glucose‐mediated GLP‐1 secretion was partially dependent on the opening of voltage dependent Ca2+ channels [190, 204]. A feature of enteroendocrine L‐cells is the ability to trigger action potentials via + + mechanisms dependent on voltage‐gated sodium channels (Na V) [186, 204]. Na V are responsible for action potential initiation and propagation in excitable cells, including nerve, + muscle, and types. Na V channels were found to be expressed in GLUTag cells [190]. However, while glucose was able to fire action potentials, glucose‐induced GLP‐1 + secretion was not abolished by the Na V blocker tetrodotoxin, hence no direct link between + Na V ‐mediated action potentials and stimulation of GLP‐1 secretion was demonstrated in this in vitro model [190]. Recent experiments done with Venus‐labeled primary L‐cells have shown that GLP‐1 secretion is tetrodotoxin dependent in native L‐cells, both under basal and under + nutrient‐stimulated conditions, indicating that GLP‐1 secretion is dependent on Na V in native mature murine L‐cells [204]. + 2+ It is noteworthy that the subunit composition of Na V and Ca v channels appear to be different between GLUTag and native L‐cells. While in GLUTag cells the Na+ current is attributed to Scn1a subunit, in native L‐cells it is most likely attributed to Scn3a. Furthermore, while in 2+ 2+ 2+ GLUTag cells the Ca flux was attributed to Ca v L‐type and N‐type Ca v channels, in native L‐ 2+ cells, it appears that the N‐type Ca v channel has no influence in GLP‐1 secretion [204]. Hence, caution should be taken when using GLUTag cells as a model for the study of Na+ and Ca2+ mediated currents. Fructose, a substrate for the facilitative fructose transporter GLUT 5, was shown to stimulate GLP‐1 secretion from GLUTag cells [191] and from perfused rat intestine [284]. Perfusion of rat ileum with fructose dissolved in distillate water increased GLP‐1 secretion by 18‐fold, compared to distillate water alone. Fructose prepared in sodium chloride induced GLP‐ 1 secretion by 10‐fold compared to control experiments infusing sodium chloride alone [284]. Therefore, it is believed that fructose stimulates GLP‐1 secretion from rat ileum in a sodium‐ independent manner. However, it is not completely understood how fructose stimulates GLP‐1 secretion. Methyl‐‐glucopyranoside (‐MDG), a non‐metabolizable sugar, promotes GLP‐1 secretion from the perfused rat ileum in a sodium‐dependent manner [284, 285]. Furthermore, in vivo

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studies demonstrated a role for non‐metabolizable sugars mediating GLP‐1 secretion. For example, sub‐chronic administration of ‐MDG resulted in a reduction of ambient glucose levels and in the glucose excursions during an oral glucose tolerance test in diabetic db/db mice. The improvement in glucose control was associated with an increase in circulating GLP‐1 [286]. ‐MDG is a specific substrate for sodium glucose transporters (SGLTs) and has no affinity for facilitative glucose transporters. SGLTs are electrogenic transporters localized in the plasma membrane and actively co‐transport glucose and sodium ions in a 1:2 ratio, therefore producing electrogenic signals [287]. ‐MGD was shown to stimulate electrical activity and promote GLP‐1 secretion in both GLUTag cells and primary L‐cell cultures [191, 193]. The effect of ‐MGD is most likely mediated by a mechanism that involves the electrogenic action of SGLTs as they couple positively charged sodium influx with sugar transport uptake. This mechanism will concentrate sugars together with positively charged Na+ in the intracellular compartment triggering action potentials, membrane depolarization, extracellular calcium uptake and GLP‐1 exocytosis. RT‐PCR analysis demonstrated the expression of the electrogenic SGLT1, SGLT3 and the facilitative glucose transporter GLUT1 and fructose transporter GLUT5 in GLUTag and native L cells [191, 193]. Interestingly and in contrast to islet ‐cells, GLUTag and native L cells do not express the facilitative sugar transporter GLUT2 [191]. Immunostaining showed SGLT1 to be localized to the apical membrane of L‐cells and enterocytes, indicating a potential role in nutrient sensing. The involvement of the electrogenic SGLTs on sugar sensing and transduction of carbohydrate‐mediated GLP‐1 secretion was demonstrated in studies where GLUTag cells were exposed to the SGLT1 and SGLT3 specific inhibitor, phlorizin. Competitive inhibition of SGLT1 and SGLT3 blocked glucose and ‐MDG mediated GLP‐1 secretion but had no effect on fructose mediated GLP‐1 secretion [191]. However, it has been reported that the human SGLT3 (hSGLT3) lacks glucose transporter capabilities and acts as a glucose‐sensitive Na+ transporter. Electrophysiological assays using the Xenopus laevis oocyte expression system, expressing hSGLT3, demonstrated that glucose caused a specific, phlorizin‐sensitive, Na+‐dependent depolarization of the membrane potential independent of glucose uptake [288].

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Sweet‐taste receptors signal transduction Another sugar sensing mechanism that has been identified in enteroendocrine cells involves the sweet‐taste receptors pathway [289]. For sweet, umami, and bitter taste, the signal transduction cascade is initiated by tastants binding to G‐protein coupled receptors (GPCRs). Sweet taste signals are mediated by the heterodimer of TAS1R2 and TAS1R3 [290‐292]; umami taste signals are mediated by TAS1R3 plus TAS1R1 and a truncated type 4 metabotropic ; and bitter taste signals are detected by the TAS2R family [293]. The functional sweet taste receptor recognizes sweet‐tasting molecules as diverse as sucrose, saccharin, indulcin and acesulfame‐K [291]. Sweet taste receptors couple through the G‐protein gustducin to AC‐generated cAMP and IP3 [294, 295]. Gustducin is a heterotrimeric G‐protein made of ‐gustducin, G3, and G13 [296]. Upon activation, heterotrimeric gustducin separates into  and  subunits. The ‐subunit of gustducin activates AC leading to an increase of cAMP [297]. The  subunit of gustducin activates phospholipase Cb2 (PLCb2). PLCb2 catalyzes the formation of IP3 leading to release of calcium from intracellular stores, activation of the ion channel Trpm5 and entry of monovalent cations [298].

Expression of components of the sweet taste receptor‐signaling cascade in L‐cells It has been reported that α‐gustducin is expressed in mouse duodenal GLP‐1‐containing cells and RT‐PCR of laser‐captured human intestinal cells immunostained for GLP‐1 confirmed co‐expression of GLP‐1 and α‐gustducin [289, 299]. Immunofluorescence studies demonstrated that ‐gustducin localized to enteroendocrine L cells that express peptide YY and glucagon‐like peptide‐1 in the human colonic mucosa [289, 300]. Expression of TAS1R2 and TAS1R3 has been demonstrated in the mouse and rat intestine [301, 302]. The transcript for human TAS1R3 (h ‐ TAS1R3) was also detected in human intestine and in the human intestinal cell lines (HuTu‐80 and NCI‐H716 cells) [300, 303].

Sweet test receptors and GLP‐1 secretion The importance of the sweet taste receptor‐signaling pathway in GLP‐1 secretion remains controversial. Studies in human NCI‐H716 enteroendocrine L‐cells demonstrated that reduction of the levels of α‐gustducin transcripts by small interfering RNA (siRNA) resulted in a decrease of glucose‐mediated GLP‐1 secretion. Furthermore, the sweetener sucralose (1mM and 5mM)

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was shown to stimulate GLP‐1 secretion in NCI‐H716 cells in a concentration‐dependent manner. Mice lacking ‐gustducin have impaired GLP‐1 secretion in response to luminal glucose when compared to littermate controls [289]. Moreover, isolated duodenum and duodenal villi from ‐gustducin null mice also showed decreased GLP‐1 secretion in response to glucose. In contrast, native purified murine L‐cells were found not to express high levels of the components of the sweet‐taste receptor pathway and artificial sweeteners such us sucralose (1mM ) did not stimulate GLP‐1 secretion and did not increase intracellular calcium from primary L‐cell cultures [193]. At higher concentration (20mM), sucralose was able to stimulate GLP‐1 secretion from primary colonic cultures, but not from duodenal cultures, an effect additive to glucose‐mediated stimulation of GLP‐1 secretion [193]. Transgenic mice expressing enhanced green fluorescent protein under the control of the ion channel Trpm5 promoter also demonstrated that the Trpm5, essential component of sweet taste receptor signaling cascade, does not co‐localize with GLP‐1 in the mouse [303]. In vivo studies in rodents and healthy individuals have failed to demonstrate an effect of artificial sweeteners in mediating GLP‐1 secretion and improving glucose control. For example, in a study designed to determine whether glucose excursions were improved and whether GIP and GLP‐1 were released in response to oral sweeteners; overnight‐fasted Wistar rats and fed Zucker diabetic fatty (ZDF) rats were administered oral sweeteners prior to intraperitoneal glucose tolerance tests (IPGTTs). The results of this study showed that artificial sweeteners did not improve glucose excursions and did not acutely enhance the release of incretin hormones in vivo. Studies in humans have also failed to show an effect of artificial sweeteners in glucose control and incretin release. For example, subchronic (3 months) dietary supplementation with sucralose (667 mg daily) did not alter glycated hemoglobin in patients with type 2 diabetes [304]. A separate study performed in healthy individuals that received, in a randomized single‐ blind fashion, an intragastric infusion of either sucralose or vehicle failed to demonstrate artificial sweetener‐mediated stimulation of GLP‐1 or GIP release [305]. Nevertheless, is important to note that a study has reported an enhancement of oral glucose‐stimulated GLP‐1 release when diet‐soda rather than carbonated water was administered before glucose [306], however the increased in circulating GLP‐1 did not translate to better glucose control. The link

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between the observed increased in circulating GLP‐1 and activation of sweet‐taste receptors was not demonstrated in this study. Bitter‐taste receptors belong to the TAS2R family and have been found to be expressed in the human colon [300]. Activation of bitter taste receptors promotes the synthesis of second messengers leading to the release of Ca2+ from intracellular stores and/or modulates the gating of ion channels that mediate Ca2+ entry into neuroepithelial taste cells [294]. In vitro studies using the STC‐1 cell line have led to the hypothesis that bitter tastans could modulate GLP‐1 secretion in vivo. In these studies denatonium (bitter taste stimulus) induced a rapid and dose‐ 2+ dependent increased of intracellular calcium [Ca ]i [307]. Denatonium is known to transduce its effect by binding to the TAS2R8. RT‐PCR analysis confirmed the presence of TAS2R8 in STC‐1 cells. Activation of TAS2R8 by denatonium couples to activation of the phospholipase C, PLCB2. When STC‐1 cells were pre‐treated with the PLC inhibitor U‐73122, denatonium‐mediated 2+ increases in intracellular calcium [Ca ]i were completely abolished [307]. In addition, denatonium was shown to stimulate CCK secretion from STC‐1 cells and this effect was 2+ dependent on L‐type voltage‐sensitive Ca v channels [308]. A recent study in mice has shown that intragastric administration of bitter tastants increased circulating in a ‐gustducin‐dependent manner and delayed gastric emptying in a GLP‐1‐ and CCK‐independent and TAS2R‐dependent manner [309]. In contrast, a study in healthy individuals where equisweet and equibitter solutions were administered intragastrically showed that both sweet and bitter tastants had no effect on the rate of gastric emptying [310]. Interestingly, a candidate gene study within the Amish Family Diabetes community assessed the association of taste receptor variants with the diagnosis of type 2 diabetes mellitus. It was found that one single nucleotide polymorphism (SNP) in TAS2R9 was associated with impaired glucose homeostasis and insulin release during OGTT [311]. There are currently no reports demonstrating that bitter tastants are able to modulate GLP‐1 secretion in humans.

Protein‐mediated GLP‐1 secretion The effect of proteins on GLP‐1 secretion has not been extensively characterized. It is believed that proteins are required to be hydrolyzed into di‐tri peptide (peptones) in order to stimulate GLP‐1 secretion [312] while intact protein appear to be less effective [206]. Meat hydrolase (MH) stimulated hormone secretion in the rat isolated colon and from the GLUTag,

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STC‐1 and NCI‐H716 cell lines via a mechanism partially dependent on voltage gated calcium channels [203, 313, 314]. Peptones enter the cell via the proton‐coupled di‐tri peptide co‐transporter PepT1/PepT2 (SLC15A1/SLC15A2). These transporters promote peptone uptake independent of Na+, but they are dependent on an acidic environment (reviewed in [315]). PepT1/PepT2 is a low affinity and high capacity transporter system with broad substrate specificity. By coupling the transport of peptones to proton (H+) influx, action potentials and membrane depolarization are triggered, which in the L‐cell has been demonstrated to be a critical event in the regulation of GLP‐1 secretion. In addition, small peptides and amino acids have been found to bind and activate a series of GPCRs on L‐cells. For example, GPR93 was proposed to transduce peptone actions and to promote CCK secretion from STC‐1 cells [313]. GPRC6A has been identified as an amino acid receptor expressed in the intestine [316], however its biological functions have not been described. Tas1R1/Tas1R3 heterodimer receptor is activated by aliphatic amino acids like the umami tastan L‐glutamate and is expressed in enteroendocrine cells [290]. However, no association between this receptor and GLP‐1 secretion has been reported to date. Studies in human NCI‐H716 cells have shown that peptones as well as essential amino acids stimulated GLP‐1 secretion signaling through a mechanism that involved the activation of mitogen‐activated protein kinase (ERK 1/2) while non‐essential amino acids had no effect [314]. For example, incubation of NCI‐H716 cells with SB203580 (a p38 MAPK inhibitor) or with the MEK1/2 inhibitor UO126 resulted in a significant reduction of MH‐mediated GLP‐1 secretion [314]. While ileal infusion of peptones rapidly elevated GLP‐1 levels in the portal effluent of rats, a mixture of amino acids were weak stimulants [279]. Hence, amino acid mixtures have been found to be less reliable stimulus [203, 207, 279]. However, studies in GLUTag cells demonstrated a role for glutamine as a potent stimulus for the regulation of GLP‐1 secretion. Moreover, human studies in lean, obese and type 2 diabetic subjects demonstrated that glutamine administration prior to ingestion of a low fat meal resulted in improved postprandial glycemia, and increased circulating levels of GLP‐1 and insulin [317, 318]. Glutamine can be

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transported into mammalian cells by several Na+‐dependent and Na+‐independent glutamine transporters systems; for review see [319]. NCI‐H716 and GLUTag cells were shown to express Na+‐dependent glutamine transporters such as ATA‐2 (system A: electrogenic, Na+‐coupled neutral amino acid transporter), ASCT‐2 (non‐electrogenic Na+ coupled amino acid transporter) and y+LAT2 (a Na+ coupled amino acid exchanger) [314]. A feature of system A‐electrogenic, Na+‐coupled neutral amino acid transporter is the ability to transport the amino acid analogue methylaminoisobutyric acid (MeAIB). MeAIB has been shown to increase intracellular calcium levels and to promote a moderate and transient increase in GLP‐1 secretion from GLUTag and primary colonic L cell cultures [189, 205]. Thus, a role for system A transporters, specially the abundantly expressed ATA‐2, in the electrogenic coupled transport of glutamine has been postulated. By this mechanism, glutamine molecules are transported into the cell coupled to Na+ ions without a counter efflux of H+ ions resulting in membrane depolarization, calcium influx and GLP‐1 secretion [320]. Studies in NCI‐H716, GLUTag and murine native L‐cells cultured under conditions commonly used to clamp a depolarized plasma membrane, exposing cells to diazoxide and KCL, demonstrated that glutamine “initiates” GLP‐1 secretion via mechanisms that do not involve K‐

ATP channels [189]. Glutamine triggers membrane depolarization and increases intracellular + calcium [Ca2 ]i in a sodium‐dependent manner. Furthermore, the glutamine mediated increase + in intracellular calcium [Ca2 ]i was dependent on extracellular calcium influx since removal of calcium from the culture buffer completely abrogated this response [189, 205]. Thus, the involvement of voltage‐dependent calcium gated channels appears to be critical in mediating the secretagogue actions of glutamine. Glutamine also amplifies GLP‐1 secretion via a mechanism downstream of membrane depolarization and calcium influx events [189, 205]. It is noteworthy to mention that in primary murine L‐cell cultures alanine and glycine triggered small and transient increases in intracellular calcium, which were not accompanied by stimulation of GLP‐1 secretion. Leucine, proline and ‐aminobutyric acid (GABA) failed to elevate intracellular calcium in primary colonic L‐cell cultures [205].

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Mechanisms underlying fat mediated GLP‐1 secretion Fat is a potent stimulus for GLP‐1 secretion [226]. Hydrolysis of ingested triglycerides to long chain free fatty acids appears to be a critical step required for the fat‐mediated regulation of GLP‐1 secretion [321, 322]. The chemical structure of the fatty acid chain is also critical.

Studies using FRIC and GLUTag cell cultures demonstrated that monounsaturated fatty acids (MUFA) of chain length of 16 carbons or more were the most potent stimulants of GLP‐1 secretion (example: oleic acid, 18:1). Conversely, fully saturated chain resulted in complete loss of fatty acid‐mediated GLP‐1 secretion stimulation [188, 323]. Studies in humans and rats confirmed the in vitro findings. MUFA administration improved glycemia control associated with increased GLP‐1 secretion [324‐328]. The molecular mechanisms involved in fatty acid‐mediated regulation of GLP‐1 secretion are diverse and complex. Nuclear receptors such as peroxisome proliferator‐activated receptors (PPARs) act as sensors of free fatty acids and regulate the expression of proteins involved in the uptake, synthesis, storage, transport and degradation of lipids maintaining homeostasis [329]. However, it is becoming evident that many of the biological actions of fatty acids are mediated via alternative mechanisms. For example, activation of the atypical protein kinase C zeta (PKCζ) has been proposed to be the mechanism by which oleic acid stimulates GLP‐1 secretion in GLUTag cells [330]. The role of PKCζ was confirmed in studies in rats where reduction of mRNA levels by ileocolonic treatment with adenoviral PKCζ siRNA resulted in a significant reduction in GLP‐1 release by intracolonic infusions of oleic acid [331]. Moreover, free fatty acids and fatty acid derivatives nowadays are recognized as important signaling molecules that mediate a variety of physiological functions via activation of G‐protein coupled receptors (GPCRs) [332, 333].

Fatty acid receptors G‐protein coupled receptors GPR40 [334], GPR41, GPR43 [193, 335, 336], GPR119 [337] and GPR120 [338, 339] are activated by either fatty acids or fatty acid derivatives. They are expressed in murine and human L and K cells [193, 335, 338, 340‐343] and activation of these receptors stimulates GLP‐1 secretion [334, 337, 338, 340‐342, 344]. Bile acids have also been shown to promote GLP‐1 secretion, via activation of the G protein coupled receptor TGR‐5,

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[345, 346]. Below, I will briefly discuss our current knowledge of the fatty acid receptors thought to regulate GLP‐1 secretion with special emphasis on GPR119, the focus of chapter 2 and 3 of this thesis. GPR40, also named 1 (FFAR1), is highly expressed in islets ‐cells, and is activated by medium to long chain C12‐C22 free fatty acids to potentiate glucose stimulated insulin secretion [343, 347]. Moreover, analysis of mice where the coding sequence of GPR40 was replaced by the reporter gene beta‐galactosidase (‐Gal), GPR40+/LacZ, allowed for the identification of GPR40 in epithelial cells throughout the intestine [334]. Double immunohistochemistry revealed co‐expression of GPR40 with both GIP and GLP‐1. In addition, GPR40 KO mice were shown to have reduced circulating insulin, GLP‐1 and GIP levels in response to acute oral fat feeding [334].

Recently, a Phase 2 clinical study of the small synthetic GPR40 specific agonist,TAK‐875, demonstrated glucose lowering effects by stimulation of glucose‐dependent insulin secretion in type 2 diabetic subjects [348]. However, the levels of GLP‐1 and the relative importance of the incretin axis to the glucose‐lowering effects of TAK‐875 were not investigated in this study. GPR120 is activated by long‐chain fatty acids. It was shown to be expressed in L‐cells [193, 338, 339] and K‐cells [342] of human and rodent intestine. GPR120 is also highly expressed in adipose tissue where it is believed to play a role in adipocyte differentiation [349]. Its expression in islet endocrine cells remains controversial [350, 351]. Furthermore, its direct role in regulating fatty acid‐mediated insulin secretion from islet ‐cells appears to be negligible [343, 352]. Conversely, several in vitro and in vivo studies demonstrated its role mediating enteroendocrine hormone secretion [338, 339, 353]. Recent studies have identified GPR120 expression in proinflammatory macrophages and have demonstrated a role for GPR120 as an omega‐3 fatty acid sensor mediating anti‐inflammatory actions [354].

Both GPR40 and GPR120 have been identified as Gαq/11 coupled receptors linked to the activation of AKT, MAPK, and/or calcium signaling in different endocrine cell types [338, 343, 355]. The involvement of GPR40 and or GPR120 in transducing fatty acid‐mediated hormone secretion from enteroendocrine cells is controversial. GPR120 knockdown by siRNA prevented

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free fatty acid stimulation of CCK secretion from STC‐1 cells, while no effect was seen when GPR40 was knocked down [353]. In contrast, a separate study where FACS sorting was used to purify duodenal CCK‐producing cells expressing green fluorescent protein (GFP) from both GPR40 KO and WT mice demonstrated that GPR40 was necessary for long chain fatty acid regulation of CCK secretion.[245]. In addition, selective GPR120 agonists promoted GLP‐1 secretion from STC‐1 cells, while selective GPR40 agonists failed to do so [356]. In contrast, studies with GLUTag cells demonstrated that GPR40 and GPR120 were not involved in GLP‐1 secretion [330, 341]. However, in the same study, it was shown that GW9508 (an agonist that activates both GPR40 and GPR120) increased GLP‐1 secretion from NCI‐H716 cells. Thus, it is possible that the contrasting data could be due to species differences. Moreover, we could hypothesize that duodenal and colonic L‐cells could exhibit differential responses to fatty acids. If true, this could explain why STC‐1 cells, a model closely resembling duodenal enteroendocrine cells respond to fatty acids and secrete GLP‐1 and CCK via GPR120 activation while GLUTag cells, a model closely resembling colonic L‐cells fail to do so. More studies are needed to test these possibilities. GPR41 (FFAR3) / GPR43 (FFAR2) are activated by short‐chain fatty acids (SCFA) such as acetate, propionate, and butyrate acid. SCFAs are products of the fermentation of dietary fibers by the . GPR41 is preferentially activated by C3‐C5 chain length fatty acids (FA) while GPR43 is by C2‐C3 chain length FA [357]. GPR41 expression has been identified by RT‐PCR analysis in human and mouse white adipose tissue, where it is believed that its activation plays a critical role in SCFA‐mediated leptin expression [358]. In this study, short‐interfering RNA (siRNA) demonstrated that targeted knockdown of GPR41 mRNA transcripts abolished propionic acid‐mediated leptin up‐regulation in primary cultures of mouse adipocytes [358]. In contrast, a separate study performed in dissociated adipocytes and in primary cultures of mouse adipose tissue found no expression of GPR41 mRNA transcripts by RT‐PCR [359]. In the same study, GPR43 was found to be highly expressed in murine adipose tissue [359]. RT‐PCR analysis of a panel of total RNA from a variety of human tissues, demonstrated the presence of GPR41 transcripts in pancreas, spleen, bone marrow , lymph nodes and small intestine [360], while northern blot analysis demonstrated that GPR43 is highly expressed in human leukocytes and spleen [360, 361]. Western blot analysis demonstrated GPR41 protein expression in the

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human colon [362] and immunohistochemistry showed that GPR41 protein localizes to enterocytes and PYY immunopositive enteroendocrine cells in the human colon [362]. Further‐ more, both receptors have been localized by immunohistochemistry on enteroendocrine L‐cells of rats and humans [336, 363]. Quantitative PCR (qPCR) analysis of mRNA isolated from FACS‐ sorted primary L‐cells expressing a modified Venus fluorescent protein under the control of the glucagon promoter [193], has shown enrichment of GPR41 and GPR43 on L‐cells compared to non‐L‐cell populations [364]. Dietary fermentable fibers, as well as SCFAs, have been shown to stimulate GLP‐1 secretion in humans [365, 366] and rodents [367]. Studies in Chinese hamster ovary (CHO‐K1) cells, transfected with expression vectors containing human GPR41 or GPR43 cDNA, demonstrated that upon SCFA‐mediated simulation, both receptors coupled to Gi/o class [360]. Moreover, activation of both receptors with propionate, resulted in phosphorylation ERK1/2 [357, 360]. In the same study, transient expression of the coding sequence of the human GPR43 gene in COS‐

7 cells showed that GPR43 activation also couples to Gq/11 and promotes accumulation of inositol phosphate [357, 360]. The role of GPR41 and GPR43 as sensors and transducers of SCFA‐mediated GLP‐1 secretion was investigated in a recent study [364]. Using primary colonic cultures the authors showed

that SCFA stimulate GLP‐1 secretion in a Gq‐dependent pathway associated with an increase in 2+ intracellular Ca . Conversely, no coupling to Gi was detected indicating that GPR41 may exhibit no role transducing SCFA‐mediated GLP‐1 secretion. However, studies done in primary colonic cultures prepared from GPR41‐/‐ and GPR43‐/‐ mice and treated with propionate and acetate demonstrated impaired SCFA‐mediated GLP‐1 secretion in both models [364]. Further‐ more, GPR43‐/‐ and to a lesser extent GPR41‐/‐ mice exhibit reduced basal and glucose‐ stimulated active GLP‐1 levels [364]. TGR5 is a G protein coupled receptor activated by bile acids. Northern blot analysis of human tissues have identified TGR5 expression in the heart, skeletal muscle, spleen, kidney, liver, stomach, small intestine, colon, and leukocytes [368]. In rodents, TGR5 is expressed in brown adipose tissue (BAT). It is also expressed in human NCI‐H716 and murine GLUTag, STC‐1 enteroendocrine cell lines and in native L cells isolated from the adult mouse [193, 345, 369].

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Bile acids were shown to bind TGR5 and increase energy expenditure in rodents via a mechanism that involves activation of hormone in brown adipose tissue (BAT) and muscle [370]. Moreover, activation of TGR5 was shown to stimulate GLP‐1 secretion from NCI‐ H716, STC‐1, and GLUTag cell lines and from primary murine intestinal cell cultures [193, 345, 369]. Transgenic mice overexpressing TGR5 and kept on a high fat diet have shown improved glucose tolerance associated with a significant increase in glucose‐stimulated GLP‐1 and insulin release [369]. Conversely, TGR5‐/‐ mice fed a high fat diet showed impaired glucose tolerance compared to control mice [369]. Pharmacological activation using the TGR5 selective agonist INT‐777, a semi‐synthetic cholic acid derivative, resulted in a potentiation of glucose‐stimulated GLP‐1 secretion in wild type mice, while it had no effect in TGR5‐/‐ mice [369]. Furthermore, INT‐777 increased levels of intracellular calcium and GLP‐1 secretion in human NCI‐H716 and murine STC‐1 cell lines [369]. This effect was dependent on adenyl cyclase activity. Studies done in isolated perfused rat colon preparation [371], and in STC‐1 [345] and GLUTag cell lines [346] demonstrated that activation of TGR5 increases cAMP production and intracellular calcium concentrations. Furthermore, mixed primary murine intestinal cultures exposed to KCl in the presence of the KATP opener diazoxide, demonstrated that bile acids stimulate GLP‐1 secretion

in a KATP channel‐independent manner [372]. Transposition of the rat bilio‐pancreatic drainage to the ileum showed that enhanced release of bile directly into the distal intestine results in a significant increase of GLP‐2, a PGDP that is co‐secreted in equimolar quantities with GLP‐1 from enteroendocrine L‐cells [372]. This result is in agreement with a study that demonstrated an increase in PYY levels (a peptide co‐ secreted with GLP‐1) following bilio‐pancreatic drainage diversion to the ileum in dogs [373]. Interestingly, obese individuals that undergo bypass surgery have been shown to exhibit improved glucose tolerance associated with an elevation of up to 20 times in the levels of GLP‐ 1. Furthermore, an increased in the levels of GLP‐2 associated with increased crypt cell proliferation was seen in rat models of gastric bypass [374]. Thus, increased bile acid delivery to the distal intestine via activation of TGR5 could be the link resulting in bypass‐associated increase of PGDP levels.

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1.3 GPR119

GPR119, also named glucose‐dependent insulinotropic receptor (GDIR), was first identified using a bioinformatics approach as an orphan G‐protein coupled receptor belonging to the class A ‐like receptor family [375].

1.3.1 GPR119 Expression Human and rodent GPR119 transcripts were originally localized in the pancreas [376, 377]. Quantitative polymerase chain reaction (qPCR) showed that GPR119 expression was enriched in the islets of Langerhans [377] compared to the entire pancreas. In situ hybridization demonstrated GPR119 transcripts localize to the islets [377]. Using a polyclonal antibody raised against a synthetic peptide corresponding to the C‐terminal region of the mouse GPR119 sequence Chu et al demonstrated that GPR119 co‐localizes to insulin producing ‐cells [377]. There has been a report where immunohistochemistry and double‐immunofluorescence studies showed co‐localization of GPR119 and pancreatic polypeptide (PP) in mouse and rat islets. GPR119 was found to be expressed by qPCR in the TC1‐6 cell line [33]. However, there is no current evidence indicating that GPR119 localizes to glucagon‐secreting ‐cells in the islets of Langerhans. GPR119 is also expressed in the stomach, duodenum, jejunum, ileum and colon of humans and rodents [340]. In situ hybridization, reverse transcriptase‐PCR (RT‐PCR) and northern blot analysis demonstrated the expression of GPR119 in NCI‐H716, FRIC, STC‐1 and GLUTag enteroendocrine L‐cell lines and in murine native L and K cells [193, 340‐342].

1.3.2 GPR119 Signal Transduction

Activation of GPR119 receptor couples to Gs and increases intracellular cAMP levels [376‐ 378]. GPR119 was amplified by RT‐PCR from human pancreas cDNA, cloned into an expression vector and was stable transfected into rat hepatoma cell line (RH7777). Using this expression system it was shown that lysophosphatidylcholine induced cAMP accumulation in a dose‐ dependent manner [376]. Furthermore, using a human embryonic kidney (HEK293)‐derived cell line stable expressing the human GPR119 (hGPR119) cDNA, it was shown that the agonists oleoylethanolamide (OEA) and PSN632408 also increased cAMP levels in a dose‐dependent

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manner [337]. Furthermore, it was shown that OEA stimulates GLP‐1 secretion in a cAMP‐PKA dependent manner and this effect is dependent on intact GPR119 signaling [341]. Conversely, inositol phosphate assays in HEK293 cells transfected with the hGPR119 showed that activation of GPR119 exhibit poor coupling efficiency to Gq [377]. Moreover, co‐transfection of HEK293 cells with hGPR119 cDNA and Gq/Gi chimera that is known to redirect Gi ‐coupled receptors to the production of inositol phosphate, demonstrated that activation of GPR119 failed to induce inositol phosphate accumulation [377]. Hence, GPR119 was shown to couple selectively to Gαs .

1.3.3 GPR119 Endogenous Ligands As previously mentioned, using RH7777 cells transfected with hGPR119, Soga et al. demonstrated that oleoyl‐lysophosphatidylcholine (18:1‐LPC), lysophosphatidylethanolamine (LPE), and lysophosphatidylinositol (LPI) induced cAMP accumulation in a dose‐dependent [376]. Conversely, no effect was seen in the native cell line indicating that LPC, LPE and LPI‐ mediated cAMP accumulation was GPR119‐dependent [376]. Furthermore, in the same study, it was shown that LPC stimulated insulin secretion from the murine pancreatic NIT‐1 β‐cell line. This effect was significantly blocked when endogenous GPR119 expression was reduced by mGPR119 siRNA [376].

Phylogenetic analysis showed that GPR119 is closely related to the (CBr) [337, 379, 380]. However, using Saccharomyces cerevisiae cells transformed with human or mouse GPR119 reporter plasmids, the fatty acid derivative (arachidonylethanolamide), natural ligand of the cannabinoid receptors, was a weak activator of GPR119 [337]. Conversely, oleoylethanolamide (OEA) was a potent activator of human GPR119 in this reporter system (EC50 of 3.2M for OEA compared to EC50 >30 for 18:1‐LPC) [337]. Chu et al, have identified oleoyldopamine (OLDA) as a potent GPR119 activator that stimulates cAMP formation in HEK cells stably transfected with the human GPR119 cDNA (OLDA EC50 of 3.2 M compared to OEA (EC50 of 4.4) [381]. Interestingly, LPC did not activate GPR119 and did not increase cAMP levels in this study. Neither OEA nor OLDA are specific ligands for GPR119. Both OEA and OLDA activate the vanilloid‐responsive transient receptor potential

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vanilloid 1 (TRVP1) ion channel [382, 383]. Furthermore, OEA also activates the peroxisome proliferator‐activated receptor alpha (PPAR) [384]. 2‐oleoyl glycerol (2OG), a product of ingested fat hydrolysis, has been proposed to be a GPR119 agonist. 2OG increased circulating levels of both GLP‐1 and GIP in humans and increased intracellular cAMP levels in CHO cells stably transfected with the human GPR119 cDNA [378]. Recently, 5‐hydroxy‐eicosapentaenoic acid (5‐HEPE), an omega‐3 unsaturated fatty acid metabolite was shown to potentiate glucose‐stimulated insulin secretion and cAMP formation in murine MIN6 insulinoma cells in a GPR119‐dependent manner [385].

1.3.4 GPR119 Insulinotropic Actions

Both GLP‐1 and GIP activate their respective Gs coupled receptors (GLP‐1R and GIPR) on the ‐cell resulting in activation of AC, increase in intracellular cAMP levels and potentiation of

GSIS [386]. Hence, the expression of GPR119, a GPCR that also couples to Gs, in pancreatic ‐ cells led to the hypothesis that direct activation of this receptor could modulate insulin secretion. The insulinotropic actions of GPR119 agonists were demonstrated in the perfused rat pancreas [376], in isolated mouse and rat islets [377, 387], in hamster insulinoma cells (HIT‐T15) [377], in rat RINm5F cells transfected with human GPR119 cDNA and in mouse insulinoma Min6 cells [385, 388]. A library of around 55,000 small‐molecule compounds was screened by Arena Pharmaceuticals for their ability to stimulate cAMP production in HEK293 and RIN‐5F cells transfected with the human GPR119 cDNA [389]. AR231453 was found to potently increase cAMP levels in the above transfected cell lines [377]. Moreover, AR231453 stimulated cAMP production in hamster HIT‐T15 insulinoma cells that endogenously express GPR119 and this effect was lost when GPR119 expression was reduced by treatment with a hamster GPR119‐ selective siRNA [377]. Importantly, AR231453 potentiated insulin release in isolated rat and mouse islets incubated in 15 mM of glucose. However, AR231453 failed to stimulate insulin secretion when islets were incubated in 5mM glucose [377]. Therefore, the insulinotropic actions of GPR119 agonists was shown to be glucose‐dependent. The specificity of AR231453 to activate GPR119 was demonstrated in vitro and in vivo. AR231453 stimulated insulin release in RIN‐5F cells stable expressing the human GPR119 receptor but not in native RIN‐5F cells, lacking

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GPR119 expression. Moreover, AR231453 had no effect on glucose‐stimulated insulin secretion in islets isolated from GPR119‐/‐ mice. In addition, AR231453 improved glucose homeostasis and increased glucose‐stimulated insulin levels during oral glucose tolerance tests (OGTT) in WT mice and in rodent models of type 2 diabetes like db/db mice and ZDF rats, but failed to do so in GPR119‐/‐ mice [377]. Interestingly, it was observed that the glucoregulatory capacity of AR231453 was reduced during an intraperitoneal glucose tolerance test (IPGTT) [377]. Furthermore, administration of the GLP‐1R antagonist exendin‐(9‐39) reduced the glucoregulatory capacity of AR231453 [340]. Conversely, administration of the DPP‐4 inhibitor sitagliptin improved AR231453‐mediated glucose control [340]. These results suggested that activation of GPR119 may promote glucose control via incretin‐mediated mechanisms. In fact, GPR119 agonists were shown to act as incretin secretagogues, and this will be discussed in the next section.

1.3.5 GPR119 Agonists and GLP‐1 Secretion

As mentioned above, GPR119 was found to be expressed in the stomach, duodenum, jejunum, ileum and colon of humans and rodents [340]; in NCI‐H716, FRIC, STC‐1 and GLUTag enteroendocrine L‐cell lines; and in murine native L and K cells [193, 340‐342]. AR231453 and PSN632408 have been shown to stimulate GLP‐1 secretion from GLUTag cells [340, 341, 390]. Furthermore, PSN632408 stimulated GLP‐1 secretion from human NCI‐H716 cells [341]. In addition, the natural ligand OEA has been shown to stimulate GLP‐1 secretion from GLUTag cells via activation of the cAMP‐PKA signaling pathway [341]. Transfecting GLUTag cells with siRNA, targeting the coding sequence of murine GPR119, demonstrated that OEA‐mediated GLP‐1 secretion was GPR119 dependent [341]. Studies in vivo confirmed the in vitro findings. Intraluminal administration of OEA to euglycemic rats significantly increased the levels of bioactive GLP‐1 compared to vehicle‐ infused rats [341]. Administration of AR231453 to WT mice increased plasma levels of active GLP‐1 and GIP and this action was lost in GPR119 ‐/‐ mice [340, 341, 390]. Recently, 5‐HEPE was shown to increase cAMP and stimulate GLP‐1 secretion from human intestinal adenocarcinoma cells (HuTu80) in a GPR119‐dependent manner [385].

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The insulinotropic effects of GPR119 agonists given during OGTT could be mediated both directly via activation of GPR119 expressed on islet β‐cells and indirectly, via the enhancement of GLP‐1 secretion. However, the mechanism(s) through which GPR119 agonists exert their glucoregulatory actions and the relative importance of for the β‐cell actions of GPR119 agonists remains incompletely understood. See Figure 1.1 for a summary of possible

mechanisms involved in GPR119 activation and the regulation of ‐cell function. We have investigated the relative contribution of direct versus indirect mechanisms involved in the improvement of glucose homeostasis following GPR119 activation. Many synthetic small molecules, specific ligands for GPR119, have been developed, reviewed in [391, 392]. In the studies presented in chapter 2, we have used AR231453 to elucidate the importance of the incretin axis in the glucoregulatory actions of GPR119 activation. Moreover, we assessed AR231453 actions on the control of gastric emptying and the involvement of gut‐ peptides candidates as mediators of these actions. In chapter 3 we have used AR881, another small synthetic specific GPR119 agonist provided by Arena Pharmaceuticals, to investigate the role of endogenous GPR119 signaling in islet β‐cells susceptibility to streptozotocin‐induced apoptosis and in β‐cell response to prolonged HF feeding in mice.

1.3.6 GPCRs and ‐cell Cytoprotection An important feature of β‐cell GPCRs coupled to cAMP generation is their ability to protect the ‐cell from external injury, glucolipotoxicity and endoplasmic reticulum (ER) stress [71, 119, 132, 393, 394]. Activation of the GLP‐1R has been shown to inhibit ‐cell death. For example, GLP‐1 improved morphology and function and reduced the rate of apoptosis of freshly isolated human islets [133]. Animal models also provided evidence for the anti‐apoptotic actions of GLP‐ 1. For example, mice treated with exendin‐4, a potent GLP‐1R agonist that is resistant to DPP‐4 inactivation, were protected against streptozotocin (STZ)‐induced ‐cell injury [132]. Furthermore, in murine models of obesity and diabetes GLP‐1R activation with exendin‐4 improved glucose control, increased ‐cell mass, proliferation, and reduced ER stress and apoptosis [119, 395]. Conversely, Glp1r‐/‐ mice are more susceptible to STZ‐induced apoptosis compared to WT controls [132, 396]. Hence, GPR119 could potentially play a role in ‐cell cytoprotection either directly or indirectly via stimulation of GLP‐1 secretion. In Chapter 3 we

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assessed whether ‐cells from mice lacking GPR119 were more susceptible to STZ‐induced apoptosis and to the deleterious actions of diet‐induced obesity. Conversely, we investigated whether GPR119 activation protects the ‐cell from cytotoxic injury.

1.3.7 GPR119 Activation and Regulation of Gastric Emptying and Food Intake GLP‐1 reduces the rate of gastric emptying, increases satiety, and reduces food intake in rodents and humans [397, 398]. Hence, it was hypothesized that GPR119 agonists could potentially exert, directly or indirectly, a hypophagic effect. Administration of the small‐ molecule GPR119 agonist PSN632408 to rats resulted in inhibition of gastric emptying and food intake [337, 399, 400] and led to a reduction in weight gain, fat mass, and plasma triglyceride levels when fed a high fat diet (HFD) [337]. However, the signaling pathways activated by PSN632408 substantially differ from those activated by the GPR119 natural ligand OEA and it was suggested that although PSN632408 activates GPR119 it might also activate GPR119‐independent pathways [388]. Furthermore, OEA belongs to the cannabinoid system that controls food intake and promotes satiety [401, 402]. However, its anorectic effects were shown to be independent of GPR119 and dependent on PPAR activation [387]. Finally, Chu et al reported that the highly specific agonist, AR231453, had no effect on food intake in mice; however, the effect of GPR119 activation on the rate of gastric emptying was not evaluated in this study [340]. In Chapter 2, we investigated the effect of acute AR231453 administration on the control of gastric emptying and the mechanisms involved.

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Agonist GPR119

K cell Agonist GPR119 Agonist GPR119  Cell GIP + AC Gs + AC ATP GIP-R + ATP c AMP GLP-1R PKA PKA cAMP EPAC

[Ca 2+] metabolism Insulin - GLP-1 L Cell Glucose K+

Ca

Figure 1.1 Proposed mechanisms for how GPR119 activation regulates ‐cell function GPR119 is expressed in islets, enteroendocrine L and K cells, and in the brain of rodents. As such, GPR119 agonists could transduce their glucoregulatory actions by direct activation of receptors present in the ‐cell, or indirectly by activation of receptors present in the L and K cells inducing incretin secretion and/or by activation of receptors present in the brain.

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1.4 NOVEL GPCRs MEDIATING GLP‐1 SECRETION We have now identified a novel signaling pathway by which progesterone regulates GLP‐1 secretion via activation of membrane‐bound progesterone receptors (studies described in chapter 4).

1.4.1 Progesterone: Genomic vs. Non‐Genomic Actions Steroid hormones are important signaling molecules regulating reproduction, differentiation, cell proliferation, apoptosis, cell remodeling, inflammation, metabolism, homeostasis and brain functions [403]. Receptors for steroid hormones are members of a super family of nuclear receptors that function as ligand‐“hormone”‐activated transcription factors [404]. The steroid hormone progesterone (P4) is the physiological ligand for the classical full length B and the N‐terminus truncated A progesterone receptor (PR) isoforms [405]. In humans, the PRA and PRB are transcribed from a single gene by alternate transcription initiation from two distinct promoters [406‐408]. Classical steroid receptors are very dynamic; they are constantly shuttled between the nucleus and the cytosol [409]. While PRA is mostly located in the nucleus, PRB distributes between the cytosol and the nucleus [410]. They are both co‐synthesized in many cell types and hormone addition shifts their localization towards the nucleus. In the absence of ligand, PRs form an inactive complex with chaperone molecules such as heat shock protein (HSP) 90, 70, 40 [411, 412]. Upon exposure to progesterone (P4), PR undergoes a conformational change, dissociates from HSPs, and dimerizes. Ligand‐bound receptor dimers then translocate to the nucleus, associate to a variety of cofactors (coactivators or corepressors), recognize cis‐acting hormone response elements (HRE) in the promoter of target genes and regulate their rate of transcription [403, 413‐416]. The process described above is known as the classical or “genomic” pathway. It requires hours to days to fully activate and integrate a response to P4. Furthermore, this pathway is sensitive to inhibitors of transcription and to inhibitors of translation. However, many of the biological actions of progesterone occur rapidly, within minutes, and are referred to as the non‐

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classical or “non‐genomic” pathway, reviewed in [417]. This process is insensitive to inhibitors of transcription and to classical progesterone receptor antagonists [418]. Progesterone‐induced non‐genomic actions have been shown to stimulate, in a tissue‐specific manner, many signaling pathways including activation of protein kinase cascades (MAPKs, PKC), inhibition of cAMP production, modulation of ion channels and increases in intracellular calcium concentration; reviewed in [419, 420]. The mechanisms involved in transducing the non‐genomic actions of progesterone are complex and not completely understood. For the purpose of this introduction, I will briefly summarize current models.

1.4.2 Non‐Receptor Mediated Non‐Genomic Progesterone Actions Steroids are lipophilic molecules that could potentially modify the lipid bilayer fluidity of the plasma membrane and alter protein signaling without interacting with a receptor. For example, P4 was proposed to rapidly block voltage gated Kv channels and to trigger membrane potential in T‐lymphocytes through this mechanism [421]. However, a supraphysiological concentration of P4 was required to induce this effect. Furthermore, the same effect was seen with the progesterone receptor antagonist RU486 [421]. Hence, the physiological relevance of this mechanism of action remains controversial.

1.4.3 Ion Channel Modulation Mediating Non‐Genomic Progesterone Actions The antiepileptic effect of P4 remains in mice lacking the classical progesterone receptors (PRs), [422]. It has been postulated that P4 protects against seizures in various animal models of epilepsy via a non‐genomic mechanism involving the modulation of the GABA receptor [423‐ 425]. In this model, P4 metabolizes to , a neurosteroid that is a potent modulator of the chloride ion channel GABAa receptor. Binding of allopregnanolone results in rapid inhibition of the GABA receptor and decreased neuronal excitability [426]. Thus, P4 anticonvulsant actions are the result of a rapid non‐genomic inhibition of neuronal excitability [426].

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1.4.4 Nuclear Receptors (PRs) Mediating Non‐Genomic Progesterone Actions One of the best‐characterized non‐genomic signaling pathways activated by steroids is the Src/ras/raf/MAP kinase pathway. Progesterone rapidly activates MAPK and it has been proposed that the transduction mechanisms involve the activation of a subset of classical PRs localized to the plasma membrane; reviewed in [419]. However, how PRs lacking defined membrane targeting motifs can associate to the plasma membrane to initiate rapid cell surface P4‐induced signaling transduction events is unclear [427]. A site for post‐translational covalent addition of fatty acids (S‐palmitoylation) has been identified in the ligand‐binding domain of human PRs [428]. Palmitoylation could potentially confer a recognition site and direct a fraction of PRs to the plasma membrane. However, it is not known if this site is conserved in other species. For example, Tian et al have not been able to detect PR palmitoylation site in Xenopus, [429]. Furthermore, in vivo, only a very small portion of PRs has been found to be associated to the plasma membrane. Another proposed mechanism has been delineated from studies of P4‐induced MAPK activation in human breast cancer cells [430]. Human nuclear progesterone receptor (hPRA/B) contains a polyproline motif in the N‐terminal domain. This motif allows the hPRA/B receptor to bind to the SH3 domain of membrane‐bound Src protein initiating the activation of the mitogen‐activated protein kinases cascade leading to ERK1/2 activation. However, this motif has not been found in other species such as mouse, Zebrafish and Xenopus. Hence, the biological relevance of the above‐described mechanism is not known in these species. Cross talk between the estrogen receptor  (ER) and PRB has been linked to P4‐induced rapid MAPK activation. It has been reported that PRB activation of ERK1/2 depends on the presence of ligand free ER. In this model, in the absence of P4, the NH2‐terminal domain of PRB interacts and forms a complex with ER in the cytoplasm. Upon P4 administration, the ligand‐bound PRB dissociates from ER. Free ER will then directly interact with the SH2 domain of Src activating it. This event will initiate a signal cascade resulting in downstream activation of ERK1/2 [431]. PRA does not appear to be able to exert this mechanism of action since its truncated NH2‐terminal domain lacks the sequence required for PR‐ER binding [431].

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1.4.5 Progesterone Receptor Membrane Component 1 and 2 (PGRMC1, PGRMC2) A putative progesterone binding protein was cloned from pig microsomal membranes and from pig vascular smooth muscle cells [432, 433]. PGRMs have been investigated mainly in the reproductive system and the kidneys. Studies have suggested a role for both PGRM1 and PGRM2 in the progesterone‐induced acrosomal reaction in mammalian sperm [434] while activation of PGRM1 exerts an anti‐apoptotic effect in granulose/luteal cells [435]. The intracellular localization of this protein remains a subject of debate. While subcellular fractionation has revealed localization to endoplasmic reticulum membranes [436], association of PGRM1 to the plasma membrane has been reported in spontaneously immortalized granulosa cells [437].

1.4.6 Novel GPCRs Mediating Non‐genomic Progesterone Actions Zhu et al constructed a cDNA library from total RNA extracted from sea trout ovarian tissue. Expressed fusion proteins were screened using a monoclonal antibody generated against a progestin involved in oocyte maturation. By this strategy, they were able to identify a new receptor, named membrane‐bound progesterone receptor (mPR) that was shown to be localized to the plasma membrane and to be selectively activated by progesterone [438]. Blast search of the database using the sea trout sequence allowed for the identification of human homologues [439]. mPR isoforms exhibit GPCR‐like structural characteristics and functions including an extracellular N‐terminus, a seven transmembrane domain, an intracellular C‐terminus and the ability to couple to heterotrimeric G‐proteins and activate a variety of non‐genomic signaling pathways, [439, 440]. Phylogenetic analysis revealed that mPRs are well conserved across a variety of vertebrate species from fish to humans [439]. Three isoforms have been identified, named mPR, mPR and mPR [439, 441]. Based on their sequence these membrane progesterone receptor isoforms have been grouped under a superfamily of receptors called the progestin and adiponectin receptor family (PAQR) [442] and have been renamed as follow: mPR (PAQR7), mPR (PAQR8), mPR (PAQR5). While PAQR7 and PAQR8 share 50% homology, PAQR5 is more divergent and probably regulates different biological processes. Northern blot analysis of human tissues and quantitative PCR of fish mRNA have shown that PAQR7 is expressed mainly in reproductive tissues and less abundantly in the bladder, and the kidney. PAQR8 is expressed in

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the brain and neurons, while PAQR5 is expressed in cells of epithelial tissues like the lungs, liver, intestine, kidney and fallopian tubes [439, 443, 444].

mPR was shown to couple to Gi in human breast carcinoma cells MDA‐MB‐231 transfected with expression vectors containing putative coding sequences of seatrout mPR [438]. In addition, progesterone (P4) treatment of plasma membrane fractions isolated from human pregnant myometrial tissue, expressing PAQR7 and PAQR8, caused rapid activation of

Gi but not Gq/11 [445]. Moreover, incubation of human myometrial membranes with P4 and with bovine serum albumin‐P4 bound conjugate (BSA‐P4), a complex that because of its size is unable to enter the intracellular compartment, resulted in a rapid and transient reduction in basal and isoproterenol‐induced cAMP levels [445]. This effect was pertussis toxin sensitive [445]. Co‐immunoprecipitation of solubilized myocyte membranes with an antibody directed against the Gαi subunit, followed by western blotting with PAQR7 and PAQR8 antibodies, demonstrated that PAQR7 and PAQR8 are coupled to Gαi in human myocytes [445].

P4 and BSA‐P4 induced a rapid increase in intracellular calcium via Gq/11–PLC activation in confluent female and male rat osteoblasts. Using antibodies against the various PLC isoforms, it

2+ was shown that only PLC‐β1 and PLC‐β3 were involved in the Ca mobilization and IP3 formation induced by the progestins in female and male osteoblast [446]. In this study, it was suggested that P4 and BSA‐P4 were directly activating membrane‐localized G‐protein receptors leading to the very rapid formation of second messengers without involving the . However, the receptors involved were not identified. Steroids stimulate ERK1/2 phosphorylation in human breast carcinoma cells MDA‐MB‐231 transfected with seatrout mPR cDNA [438]. However, treatment of human myometrial cells with P4 and BSA‐P4 failed to increase ERK1/2 phosphorylation while they stimulated P38 MAPK activation [445]. Western blot analysis and RT‐PCR studies demonstrated mRNA and protein co‐ expression of PAQR5 and PAQR7 in the proximal tubules of kidneys of male and female mice. As with human myometrial cells, P4 failed to stimulate ERK1/2 phosphorylation in dissected proximal tubules of murine kidneys that were perfused with the steroid [447]. Despite the restricted tissue specific expression of PAQR5, PAQR7, and PAQR8 isoforms, there are currently few studies looking at the biological functions of mPRs in tissues other than the reproductive system. In addition, there is evidence for the possibility of crosstalk between

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mPR and PR. For example, there is a report where human myometrial cells were transfected with the PR‐B isoform, in order to shift the PR‐A/PR‐B ratio in favor of a functional nuclear receptor. In this system, P4 induced transactivation of a luciferase reporter expression vector of the glucocorticoid‐responsive element. However, this effect was reduced when cells were pre‐ treated with pertussis toxin, suggesting that, at least in human myometrial cells, activation of

Gαi‐mediated signaling potentiate P4‐activated PR [445]. Further studies are needed to investigate the possible crosstalk between mPRs and between mPRs and the classical PR.

1.5 RATIONALE FOR THE STUDIES DESCRIBED IN MY THESIS

1.5.1 Chapter 2 GPR119 is expressed in islet ‐cells where its activation stimulates cAMP accumulation and glucose‐dependent insulin secretion [376, 377, 388]. Furthermore, it is expressed in enteroendocrine L and K cells where its activation stimulates GLP‐1 and GIP release [340, 341, 381]. GPR119 agonists improve glucose homeostasis. However, the mechanism(s) involved and the relative importance of incretins for the glucoregulatory actions of GPR119 agonists remain incompletely understood. Principal Hypothesis: The incretin receptors GLP‐1 and GIP represent the dominant mechanisms for transduction of the glucoregulatory actions of GPR119 agonists. In Chapter 2 we carried out experiments to test this hypothesis. Aim 1: Pharmacological activation of GPR119: To determine whether the effects of GPR119 activation were dependent on intact incretin signaling we used genetic models of incretin receptor ablation (Glp1r‐/‐, Gipr‐/‐ and double incretin receptor knockout (DIRKO) mice) and a highly selective GPR119 agonist (AR231453). A considerable body of experimental evidence in rodents, pigs and human studies demonstrates that GLP‐1 regulates gastrointestinal motility and gastric secretion [448, 449]. Furthermore, the glucoregulatory actions of GLP‐1 are known to be mediated by a combination of complementary effects. Apart from its direct insulinotropic action in the islet ‐cell, GLP‐1 controls glucose homeostasis via inhibition of gastric emptying resulting in reduced postprandial glucose excursions [141]. Moreover, in diabetic patients, GLP‐1 still is able to

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reduce glucose excursions even in the absence of an increase in circulating insulin by slowing the transit and absorption of nutrients [142]. OEA, natural ligand for GPR119 and PPAR inhibits gastric emptying and intestinal motility in a PPAR‐independent manner [450]. Thus, it may be possible that GPR119 activation will control glucose homeostasis via multiple mechanisms including reducing the rate of gastric emptying. Secondary Hypothesis: GPR119 activation indirectly reduces the rate of gastric emptying via stimulation of GLP‐1 secretion and this mechanism contributes, at least in part, to GPR119‐ dependent glucoregulation. Aim 2: GPR119 activation and its effects on gastric emptying: To determine whether activation of GPR119 with a specific agonist reduces gastric emptying and to delineate the mechanisms involved we performed the acetaminophen absorption test in WT, GPR119‐/‐, Glp1r‐/‐, Glp2r‐/‐ mice. Furthermore, GPR119 agonists have been shown to stimulate PYY release from isolated gut mucosa [451] and PYY inhibits the rate of gastric emptying [452]; thus, we also investigated whether GPR119 activation reduces the rate of gastric emptying in WT mice pre‐treated with the PYY receptor (Y2R) inhibitor BII0246.

1.5.2 Chapter 3 Hypothesis: Endogenous GPR119 signaling is critical for β‐cell survival and GPR119 agonists protect the β‐cell directly or indirectly through stimulation of GLP‐1 and GIP secretion. Aim 1 Gain of function: Evaluate the role of pharmacological activation of GPR119 in ‐cell cytoprotection against STZ‐induced injury. We determined whether sub‐chronic administration of the GPR119 agonist AR881, protects β‐cells from STZ‐induced apoptosis in C57BL/6 mice. Aim 2 Loss of function: Evaluate ‐cell susceptibility to STZ‐induced apoptosis in GPR119‐/‐ mice. To determine whether the loss of GPR119 enhances the susceptibility of the β‐cell to external injury we have assessed the degree of apoptosis in GPR119‐/‐ and littermate control +/+ mice using the low dose streptozotocin model of ‐cell injury [132]. Aim 3 Loss of function: Evaluate ‐cell susceptibility to lipotoxicity in GPR119‐/‐ mice fed a high fat diet. To determine whether ablation of GPR119 signaling results in enhanced ‐cell susceptibility to the deleterious effects of lipids we assessed ‐cell gene expression, ‐cell

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function and ‐cell adaptation in GPR119‐/‐ and littermate controls fed regular chow (RC) or high fat diet (HFD). The results of these studies are described in Chapter 3 of this thesis.

1.5.3 Chapter 4 GLP‐1 and glucagon exert opposite effects on glucose homeostasis. GLP‐1 lowers blood glucose levels, stimulates glucose‐dependent insulin secretion and insulin biosynthesis and inhibits glucagon release; reviewed in [386, 453]. Conversely, glucagon is the major counter‐ regulatory hormone to insulin. It is important in the maintenance of normal glucose levels, especially during fasting. Glucagon promotes glycogenolysis and gluconeogenesis while inhibiting glycolysis in the liver; reviewed in [454]. Release of GLP‐1 from L‐cells is stimulated by food ingestion and insulin. Conversely, glucagon secretion from islet α‐cells is stimulated in response to a fall in ambient plasma glucose (hypoglycemia) and inhibited, in the context of whole islets, by hyperglycemia and insulin. However, how PGDP secretion from α‐cells and L‐ cells is regulated is not completely understood. Type 2 diabetes is characterized by inappropriate regulation of hepatic glucose production, which is due, at least in part, to an imbalance in the bihormonal relationship between plasma levels of glucagon and insulin. Plasma glucagon concentrations fail to decrease appropriately or, paradoxically, may even increase, after oral glucose or carbohydrate ingestion [455‐459]. Moreover, type 2 diabetic patients exhibit a reduced incretin effect in part due to reduced GLP‐ 1 secretion [12‐14]. Understanding the mechanisms that differentially regulates proglucagon gene transcription and secretion in the enteroendocrine L‐cell versus the islet ‐cell is of importance and may help to elucidate the reason behind the defective response to nutrient that is seen in type 2 diabetes. Hypothesis: Proglucagon gene expression is differentially regulated in islet ‐cells versus enteroendocrine L‐cells by a unique network of transcription factors differentially expressed in ‐cells vs. L‐cells. Aim 1: To delineate gene expression profiles by microarray technology and to identify differentially expressed transcription factors by functional network analysis in murine gut GLUTag and islet αTC1 cell lines. In Aim1, we identified that the nuclear transcription factor, progesterone receptor (PR), is

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expressed in GLUTag cells but not in α‐TC1 cells. Secondary Hypothesis: PR transactivates proglucagon gene transcription and regulates GLP‐1 secretion in GLUTag cells. Aim 2: Investigate the role of progesterone receptor in the regulation of proglucagon gene transcription and GLP‐1 secretion and delineate the signaling pathways involved. The results of these studies are presented in Chapter 4.

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Chapter 2

GPR119 REGULATES MURINE GLUCOSE HOMEOSTASIS THROUGH INCRETIN RECEPTOR‐DEPENDENT AND INDEPENDENT MECHANISMS

The work presented in this chapter has been modified from the following publication: Grace Flock, Dianne Holland and Daniel. J. Drucker. Endocrinology. 2011 Feb;152(2):374‐83.

Author contributions: Dianne Holland contributed to animal experiments, providing technical assistance (Figures: 2.1 and 2.3)

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2.1 ABSTRACT

GPR119 was originally identified as a ‐cell receptor however; GPR119 activation also promotes incretin secretion and enhances PYY action. We examined whether GPR119‐ dependent control of glucose homeostasis requires preservation of peptidergic pathways in vivo. Insulin secretion was assessed directly in islets and glucoregulation was examined in wild type (WT), single incretin receptor null, and dual incretin receptor knockout (DIRKO) mice. Experimental endpoints included plasma glucose, insulin, glucagon, GLP‐1, GIP and PYY. Gastric emptying was assessed in WT, Glp1r‐/‐, DIRKO, Glp2r‐/‐ and GPR119‐/‐ mice treated with the GPR119 agonist AR231453. AR231453 stimulated insulin secretion from WT and DIRKO islets in a glucose‐dependent manner, improved glucose homeostasis and augmented plasma levels of GLP‐1, GIP and insulin in WT and Gipr‐/‐mice. In contrast, although AR231453 increased levels of GLP‐1, GIP, and insulin, it failed to lower glucose in Glp1r‐/‐ and DIRKO mice. Furthermore, AR231453 did not improve intraperitoneal glucose tolerance and had no effect on insulin action in WT and DIRKO mice. Acute GPR119 activation with AR231453 inhibited gastric emptying in Glp1r‐/‐, DIRKO, Glp2r‐/‐ and WT mice independent of the Y2R; however, AR231453 did not control gastric emptying in GPR119‐/‐ mice. Our findings demonstrate that GPR119 activation directly stimulates insulin secretion from islets in vitro, yet requires intact incretin receptor signaling and enteral glucose exposure for optimal control of glucose tolerance in vivo. In contrast, AR231453 inhibits gastric emptying independent of incretin, Y2R or Glp2 receptors through GPR119‐dependent pathways. Hence, GPR119 engages multiple complementary pathways for control of glucose homeostasis.

2.2 INTRODUCTION Enteroendocrine cells and islet ‐cells share a number of physiological regulatory mechanisms. Most notably, basal hormone secretion is maintained at a constant but low rate in the absence of nutrient ingestion; however, food intake rapidly augments hormone secretion. Enteroendocrine cells communicate with and amplify ‐cell function through secretion of peptide hormones. For example, glucose‐dependent insulinotropic peptide (GIP) secreted by K cells in the duodenum and proximal jejunum, acts as a potent incretin to enhance glucose‐

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dependent insulin secretion [460]. Similarly, a second incretin, glucagon‐like peptide‐1 (GLP‐1), secreted from gut L cells, exerts pleiotropic actions on islet ‐cells, including stimulation of insulin biosynthesis and secretion and cell proliferation, promotion of cell survival, and restoration of ‐cell glucose sensing [461]. The ability of incretin hormones to enhance ‐cell survival, restore glucose sensing to poorly responsive diabetic ‐cells, and control glycemia in a glucose‐dependent manner has fostered the development of drugs that potentiate incretin action [462]. Two new classes of therapeutic agents based on incretin action, GLP‐1R agonists, and dipeptidyl peptidase‐4 (DPP‐ 4) inhibitors, are now used to treat type 2 diabetes, [461]. The success of these agents has engendered interest in discovery and characterization of ‐cell G protein coupled receptors (GPCRs) that function in an incretin receptor‐like manner. A large number of GPCRs expressed in islet ‐cells are coupled to stimulation of glucose‐ dependent insulin secretion [463]. One of these receptors, designated GPR119 or glucose‐ dependent insulinotropic receptor (GDIR), has been independently isolated by multiple groups and characterized in different species [337, 375, 377]. GPR119 is expressed in rodent islets and β‐cell lines and GPR119 activation enhances cyclic AMP formation and stimulation of insulin secretion [341, 376, 381, 388, 464]. These findings established the initial concept that GPR119, expressed on islet ‐cells, may be a promising target for diabetes drug development through mechanisms involving direct potentiation of insulin secretion [337]. Recent findings have expanded the spectrum of GPR119 action to encompass stimulation of gut hormone secretion. The synthetic GPR119 agonist AR231453 [389] increased circulating levels of GLP‐1 and GIP in WT mice, but not in GPR119‐/‐ mice [340]. Moreover, GPR119 expression has been detected in normal L and K cells [193, 342] and in the GLUTag enteroendocrine L cell line [193, 340], and direct GPR119 activation in L cells stimulates cyclic AMP formation and GLP‐1 secretion [340, 341]. Furthermore, GPR119‐/‐ mice exhibit attenuated nutrient‐stimulated GLP‐1 release [387]. Hence, these observations imply that GPR119 activation may control glucose through multiple mechanisms, including direct activation of the ‐cell GPR, or indirectly, through potentiation of incretin secretion. Consistent with this hypothesis, partial blockade of GLP‐1R signaling with exendin (9‐39) attenuates GPR119‐dependent improvement in oral glucose tolerance in normal mice [340].

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More recent evidence has deepened the complexity of GPR119 mechanisms of action by implicating an essential role for PYY as a downstream target required for the gastrointestinal and glucoregulatory actions of GPR119 [451]. Accordingly, to determine the relative importance of gut hormones for the glucoregulatory and gastrointestinal actions of GPR119, we examined GPR119 activation using a selective GPR119 agonist in normal mice, isolated islets, and in mice with inactivation of gut hormone receptors. We also defined a potent effect of and specific pathways required for GPR119‐dependent inhibition of gastric emptying. Our findings invoke a role for both direct ‐cell action and the incretin axis in the transduction of glucoregulatory mechanisms pursuant to GPR119 activation.

2.3 RESEARCH DESIGN AND METHODS 2.3.1 Animal experiments Animal experiments were carried out according to protocols approved by the Mt. Sinai Hospital and the Toronto Centre for Phenogenomics (TCP) Animal Care Committees. Male, age matched, C57BL/6 mice were purchased from Taconic and were allowed to acclimatize to the animal facility for 1 wk before experimentation. DIRKO, Glp2r‐/‐, Gipr‐/‐, and Glp1r‐/‐ and littermate control male mice were on a C57BL/6 genetic background [465, 466] . Mice were maintained on a 12‐h light, 12‐h dark cycle. GPR119‐/‐ mice were obtained from Arena Pharmaceuticals, San Diego, CA [377].

2.3.2 Glucose tolerance tests Mice were fasted overnight, weighed and fasting blood glucose levels were measured by tail vein sampling using a glucometer (Bayer, Toronto, ON). A single oral dose of 20 mg/kg AR231453 (GPR119 agonist, Arena Pharmaceuticals) or vehicle (80% PEG400, 10% Tween 80, 10% ethanol) was given 30 min prior to oral or intraperitoneal (IP) glucose (1.5 g/kg body weight). Blood glucose levels were measured by sampling from the tail vein of gently held conscious mice, from 5‐120 min after glucose administration. At the 5‐min time point, a blood sample (150 µl) was collected and immediately mixed with 15 µl of a chilled solution containing 5000 kIU/ml Trasylol (Bayer, Toronto, ON), 32 mM EDTA, and 0.01 mM Diprotin A (Sigma St. Louis, MO) and kept on ice for assessment of total GLP‐1, total GIP, insulin and PYY levels in

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plasma. At the 15 min time point, a second blood sample (50 μl) was collected from each mouse in EDTA coated tubes (Sarstedt, Montreal, QC) for measurement of glucagon. Plasma was obtained by centrifugation at 4 C and stored at –80 C until determination of insulin (ultrasensitive mouse insulin ELISA, Alpco Diagnostics, Salem, NH), total GLP‐1 (mouse/rat total GLP‐1 assay kit Mesoscale Discovery, Gaithersburg, MD), total GIP (rat/mouse GIP (total) ELISA kit Millipore, Billerica, MA), PYY [467] (ELISA kit Alpco Diagnostics, Salem, NH) and glucagon (Millipore, Billerica, MA) levels.

2.3.3 Insulin Tolerance Test Ten‐week‐old age‐matched C57BL/6 and DIRKO male mice were fasted for 5 hrs. A single dose of 1.2 U / kg of insulin (Humulin R, Eli Lilly, On. Canada) was administered by ip injection and blood glucose was determined at 0, 15, 30, 60 and 90 minutes.

2.3.4 Arginine stimulation test Ten‐week‐old age matched C57BL/6 and DIRKO male mice were fasted overnight. A fasting blood sample was collected and mice were given a single ip dose of arginine (Arg) 1 mg/g (Sigma, St Louis, MO). A second blood sample was collected 5 min after Arg administration. Insulin was measured at fasting and 5 min after Arg administration (ultrasensitive mouse insulin

ELISA, Alpco Diagnostics, Salem, NH).

2.3.5 Insulin secretion in mouse islets Islets were prepared as described [396]. After 2 hours of incubation at 37 C, islets were handpicked into fresh medium and allowed to recover overnight. Islets with preserved architectural integrity were used for insulin secretion experiments. Batches of 10 islets were distributed into tubes, in triplicates, containing 0.5 ml Krebs‐Ringer buffer containing 2.8 or 16.7 mM glucose, with or without vehicle (Dimethyl Sulfoxide, DMSO), or the GPR119 agonist AR231453, a 300 nM solution prepared in DMSO (Arena Pharmaceuticals, San Diego, CA), exendin‐4 (10 nM), forskolin (10 uM), pituitary adenylate cyclase activating polypeptide (PACAP) (10 nM, Sigma). After incubation for 1 h at 37 C, medium was collected and stored at – 20 C for assessment of insulin secretion. Islet insulin content was extracted by transfer of islets

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to cold acid‐ethanol solution (70% ethanol, 0.18M HCl). Extracts were briefly sonicated (10 sec), and insulin was measured by RIA (Linco rat insulin RIA # RI‐13K).

2.3.6 Analysis of GPR119 expression in islets First‐strand complementary DNA was synthesized from total islet RNA using the SuperScript III reverse transcriptase synthesis system (Invitrogen, Carlsbad, CA) and random hexamers. Real‐time polymerase chain reaction was performed with the ABI Prism 7900 Sequence Detection System using TaqMan Gene Expression Assay GPR119 (Mm00731497_s1), TaqMan Universal PCR Master Mix (Applied Biosystems, Foster City, CA). IRS‐2 (Mm03038438_m1) was used as a control and levels of messenger RNA (mRNA) transcripts were normalized to levels of 18S RNA in the same samples (Hs99999901_s1).

2.3.7 Gastric emptying Liquid‐phase gastric emptying was assessed using the acetaminophen absorption test [468, 469]. C57BL/6, Glp1r‐/‐, DIRKO, GPR119‐/‐ and Glp2r‐/‐ male mice, 10‐12 weeks of age were fasted overnight and given a single dose of either AR231453 (20 mg/kg) or vehicle (80% PEG400, 10% Tween 80, 10% ethanol) 30 min before oral administration of a solution of glucose 15% and acetaminophen 1% (Sigma, St Louis, MO ) at a dose of 1.5 g/kg glucose ‐ 0.1 g/kg acetaminophen. Exendin‐4 (1 ug, ip) was used as a positive control to demonstrate inhibition of gastric emptying. Tail vein blood (50 µl) was collected into heparinized tubes at 15 and 30 min after glucose/acetaminophen administration. Plasma was separated by centrifugation at 4 C and stored at –20 C until measurement of acetaminophen levels using an enzymatic‐ spectrophotometric assay (Diagnostic Chemicals Ltd., Oxford, CT).

2.3.8 Statistical Analysis Results are presented as mean ± SEM. Statistical significance was determined using analysis of variance with Bonferroni post hoc tests or Student’s t test (as appropriate) using GraphPad Prism 4.0 (GraphPad Software Inc, La Jolla, CA). Statistical significance was noted when P < 0.05.

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2.4 RESULTS GPR119 activation and glucose tolerance As plasma levels of GLP‐1 and GIP increase following GPR119 activation [340], we assessed whether complete loss of incretin action diminishes the glucoregulatory actions of GPR119. The GPR119 agonist AR231453 and oral glucose were administered to WT, Gipr‐/‐, Glp1r‐/‐ and DIRKO mice (Figure 2.1). AR231453 significantly reduced glucose excursions following oral glucose loading in WT and Gipr‐/‐ mice (Figure 2.1A, B). In contrast, a detectable but non‐ significant reduction in glycemic excursion was observed following oral glucose loading and AR231453 treatment of Glp1r‐/‐ and DIRKO mice (Figure 2.1C, D). To elucidate the importance of enteral vs. parenteral glucose exposure for the glucoregulatory actions of GPR119 agonists, we tested whether AR231453 improved intraperitoneal glucose tolerance. Oral AR231453 administration failed to control glucose excursions in WT mice following intraperitoneal glucose challenge (Figure 2.1E). Plasma hormones assessed following intraperitoneal glucose challenge revealed an increase in GLP‐1, but no significant changes in circulating GIP, insulin or glucagon (Figure 2.2). Hence, GPR119 activation requires GLP‐1R signaling for optimal control of oral glucose tolerance and GPR119 activation in the absence of enteral glucose exposure is not sufficient to improve glucose homeostasis.

GPR119, incretin and islet hormones and insulin tolerance To interpret the findings observed with WT and knockout mice in Figure 2.1, we assessed whether AR231453 regulates circulating levels of incretin and islet hormones to a similar extent in WT and incretin receptor knockout mice. Plasma levels of total GIP were modestly increased by AR231453 in WT and Gipr‐/‐ mice (Figure 2.3A) and significantly increased in Glp1r‐/‐ and DIRKO mice (Figure 2.3A). In contrast, plasma levels of total GLP‐1 were significantly increased by AR231453 in WT, Gipr‐/‐, Glp1r‐/‐ and DIRKO mice (Figure 2.3B). Similarly, plasma insulin levels were significantly increased in WT, Gipr‐/‐, and Glp1r‐/‐ mice following oral glucose and

AR231453 administration (Figure 2.3C).

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Measure blood glucose

Tail Tail bleed bleed (150) ul (50) ul - 30 0 5 15 30 60 90 120 Time (min)

Oral Glucose (1.5 g/kg) Measure: Measure: AR2311453 Oral (panels A - C) , Total GLP1 Glucagon (20 mg/kg) IP (panel E) Total GIP Insulin

A B 20 WT 20 Gipr-/- * * Vehicle 15 15 AR 231453 10 10 400 * 400 300 ** Vehicle 300 AR231453 200

5 5 AUC 200 Glucose (mM) Glucose Glucose (mM) Glucose

AUC 100 100 (mM / min) (mM / min) (mM / 0 0 0 0 0 15 30 45 60 75 90 105120 0 15 30 45 60 75 90 105 120 Time (min) Time (min) C D 20 Glp1r-/- 25 DIRKO

15 20 15

10 400 750

300 10 500 5 200 AUC Glucose (mM) Glucose AUC Glucose (mM) 5 250 100 (mMmin) / (mM / (mM min) 0 0 0 0 0 15 30 45 60 75 90 105 120 0 15 30 45 60 75 90 105 120 Time (min) Time (min) E 25 WT 20 15 10

Glucose (mM) Glucose 5 0 0 15 30 45 60 75 90 105120 Time (min)

Figure 2.1 GPR119 activation and control of oral glucose in WT and incretin receptor knockout mice. Age‐matched male mice were fasted overnight and AR231453 (20 mg/kg) or vehicle was administered orally 30 min before an oral glucose load (OGTT) (panels A‐D) or intraperitoneal glucose load (IPGTT) (panel E) (1.5 g/kg). Blood was collected from the tail vein at 5 minutes (150 ul) and at 15 minutes (50 ul) after glucose administration. Blood glucose values and area under the curve analysis during OGTT for (A) WT, (B) Gip1r‐/‐ (C) Glp1r‐/‐, and (D) DIRKO mice treated with or without AR231453. Blood glucose values during IPGTT for WT mice treated with or without AR231453 (E) Statistical analysis was assessed by ANOVA. * p<0.05, ** p<0.01. For vehicle‐treated mice, n=12 (A), 8 (B), 10 (C) and 12 (D); for AR231453‐treated mice, n = 11 (A), 8 (B), 10 (C), 12 (D) and 10‐12 (E).

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Measure blood glucose

Tail Tail bleed bleed (150) ul (50) ul - 30 0 5 15 30 60 90 120 Time (min)

Oral ip Glucose Measure: Measure: AR2311453 (1.5 g/kg) Total GLP1 Glucagon (20 mg/kg) Total GIP Insulin

A C 0.6 60 Vehicle 0.5 50 AR231453 40 0.4 30 0.3 20 0.2 0.1

10 Insulin (ng/ml) Total GIP (pg/ml) GIP Total 0 0.0

B D 70 * 30 60 50 20 40 30 10 20 10 Glucagon (pg/ml) Total GLP1 (pg/ml) 0 0

Figure 2.2 GPR119 activation and plasma levels of GIP, GLP‐1, insulin, and glucagon in WT mice during an intraperitoneal glucose tolerance test (IPGTT). Age matched WT mice were fasted overnight and oral AR231453 (20 mg/kg) or vehicle was administered 30 min before ip glucose load (OGTT) (1.5g/kg). Plasma was obtained 5 minutes after glucose administration. This sample was used to simultaneously measure the levels of total GIP immunoreactivity (A), total GLP‐1 immunoreactivity (B) and insulin (C). A second plasma sample was collected from each mouse at 15 min after glucose for the measurement of glucagon levels (D). Statistical analysis was assessed by ANOVA. *p<0.05.

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Measure blood glucose

Tail Tail bleed bleed (150) ul (50) ul - 30 0 5 15 30 60 90 120 Time (min)

Oral Oral Glucose Measure: Measure: AR2311453 (1.5 g/kg) Total GLP1 Glucagon (20 mg/kg) Total GIP Insulin

** A *** C 350 Vehicle 0.9 ** 300 AR231453 ** 0.8 ** 0.7 250 0.6 ** 200 0.5 150 0.4 ** 100 0.3 Insulin (ng/ml) Insulin

Total GIP(pg/ml) 0.2 50 0.1 0 0.0 WT Gipr-/- Glp1r-/- DIRKO WT Gipr-/- Glp1r-/- DIRKO (n=12) (n=8) (n=11) (n=12) (n=12) (n=8) (n=11) (n=12)

B * D * 250 *** 45 * 40 * 200 35 ** ** 30 150 25 *** 20 100 ** 15 10 50 Glucagon (pg/ml) Total GLP1 (pg/ml) GLP1 Total 5 0 0 WT Gipr-/- Glp1r-/- DIRKO WT Gipr-/- Glp1r-/- DIRKO (n=12) (n=8) (n=11) (n=12) (n=12) (n=8) (n=11) (n=12)

Figure 2.3 GPR119 activation and plasma levels of GIP, GLP‐1, insulin and glucagon in WT and incretin receptor knockout mice. Age matched male mice were fasted overnight and oral AR231453 (20mg/kg) or vehicle was administered 30 minutes before an oral glucose load (OGTT). Plasma was obtained 5 minutes after oral glucose administration This sample was used to simultaneously measure the levels of total GIP immunoreactivity (A), total GLP‐1 immunoreactivity (B), insulin (C). A second plasma sample was collected from each mouse at 15 minutes after glucose for the measurement of glucagon levels (D) in WT, Gipr‐/‐, Glp1r‐/‐, and DIRKO mice. Statistical analysis was assessed by ANOVA. *p<0.05; ** p<0.01; *** p<0.001.

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Although insulin levels were significantly lower in DIRKO compared to WT mice following oral glucose alone reflecting defective β‐cell function in DIRKO islets [465, 470, 471], AR231453 significantly increased plasma insulin levels in DIRKO mice, albeit to levels lower than those observed in WT or single incretin receptor knockout mice (Figure 2.3C). AR231453 had no effect on plasma glucagon levels in WT mice (Figure 2.3D). However basal glucagon levels were modestly lower in incretin receptor knockout mice and increased significantly following oral glucose and AR231453 administration in Gipr‐/‐, Glp1r‐/‐ and DIRKO mice (Figure 2.3D). Insulin sensitivity assessed by insulin tolerance testing was comparable in WT and DIRKO mice in the presence or absence of AR231453 (Figure 2.4). Hence, GPR119 activation significantly increases GLP‐1 levels in both single and double incretin receptor knockout mice and acute GPR119 activation increases plasma insulin levels despite the complete absence of incretin receptor action.

GPR119 directly increases insulin secretion from isolated islets We next assessed whether the diminished response of DIRKO mice to AR231453 reflects a selective inability of murine ‐cells to respond to direct GPR119 activation and/or a generalized defect in secretagogue‐regulated insulin secretion from DIRKO β‐cells [465, 470, 471]. Acute administration of arginine significantly increased plasma insulin levels in both WT and DIRKO mice (Figure 2.5A), demonstrating that DIRKO mice retain the ability to robustly respond to

non‐GPCR‐mediated amino acid insulin secretagogues. One possibility that might explain altered glucoregulatory responses to AR231453 could be a change in the relative level of GPR119 expression in single incretin receptor or DIRKO islets. However, no significant differences in the relative levels of GPR119 mRNA transcripts were observed in RNA from WT, Glp1r‐/‐, Gipr‐/‐ vs. DIRKO islets (Figure 2.5B). As a control for assessment of gene expression in murine islets, we examined Irs‐2. In keeping with previous findings [396], levels of Irs‐2 mRNA transcripts were significantly reduced in islet RNA from Glp1r‐/‐ relative to Gipr‐/‐ or WT mice (Figure 2.5B). Similarly, consistent with findings in the Glp1r‐/‐ mice, Irs‐2 mRNA transcripts were also reduced in RNA from DIRKO islets (Figure 2.5B).

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5 hrs fasting Measure blood glucose

Time (min) - 30 0 90

Oral Insulin (1.2 IU/kg) AR2311453 or vehicle

AB 15 DIRKO 12 WT

Vehicle 9 AR 231453 10

6

5 3 Blood GlucoseBlood (mM) Blood Glucose (mM) Glucose Blood

0 0 0 15 30 45 60 75 90 0 15 30 45 60 75 90

Time (min) Time (min)

Figure 2.4 GPR119 activation does not modify glucose profiles in an insulin tolerance test (ITT) in WT and DIRKO mice. WT (n=10) and DIRKO (n=8) age matched mice were fasted for 5 hrs and oral AR231453 (1.2 mg/kg) or vehicle was administered 30 min before intraperitoneal (IP) insulin administration (1.2U / kg). Blood glucose values during ITT for WT (A) and DIRKO (B) mice.

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A B

WT-Vehicle (n=8) WT-Arg (n=8) * DIRKO-Vehicle (n=8) 0.45 ** 1.00 DIRKO-Arg (n=8) * 0.40 C57Bl6 n=5 0.35 Gipr -/- n=6 0.75 * 0.30 Glp1r -/- n=6 0.25 DIRKO n=5 0.50 0.20

RNA / 18s / RNA 0.15

Insulin (ng/ml) 0.25 0.10 0.05 0.00 0.00 Fasting 5 min after Arg Gpr119 Irs2

Fasting After Arg. tail bleed tail bleed

C LG-Vehicle 0 5 HG-Exendin4 HG-vehicle HG-AR231453 HG-PACAP Arginine (IP) ** 0.008 *** * 0.007 p>0.05 0.006 *** 0.005 0.004 0.003 * 0.002 0.001 0.000 C57Bl6 DIRKO insulin secreted / islet insulin content insulin / islet secreted insulin

Figure 2.5 GPR119 activation increases insulin secretion from WT and DIRKO islets. (A) Arginine stimulation increases plasma insulin levels in C57BL/6 and DIRKO mice in vivo. Mice were fasted overnight, plasma was collected to assess fasting insulin levels and a single intraperitoneal dose of either arginine (Arg) (1 mg/g) or vehicle was administered at time zero. Blood was collected from the tail vein 5 minutes later for determination of plasma insulin levels. (B) GPR119 and Irs2 gene expression in WT and incretin receptor knockout islets. GPR119 and Irs2 mRNA levels were measured by real‐time polymerase chain reaction in islets isolated from age‐matched WT, Gipr‐/‐, GLP‐1r ‐/‐, and DIRKO mice and normalized to levels of 18S RNA in the same samples. (C) Insulin secretion from WT and DIRKO islets. Islets were isolated from 10 week old C57L/6 and DIRKO mice and incubated under conditions of low glucose (LG) (2.5 mM) for 1 hour. Islets were then incubated in low glucose (LG) or high glucose (HG) (16.6 mM) and treated for 1 hr with either vehicle (DMSO), PACAP, exendin‐4 , or AR231453 . Statistical analysis was assessed by ANOVA. * p<0.05, ** p<0.01, *** p<0.001.

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We next investigated whether AR231453 directly stimulates insulin secretion from cultured murine islets in vitro. Glucose and PACAP significantly increased insulin secretion in both WT and DIRKO islets (Figure 2.5C), whereas the GLP‐1R agonist exendin‐4 increased insulin secretion in WT but not in DIRKO islets (Figure 2.5C). AR231453 significantly increased insulin secretion in a glucose‐dependent manner in both WT and DIRKO islets (Figure 2.5C). Hence, these observations demonstrate that the diminished glucoregulatory response of DIRKO mice to GPR119 activation in vivo is not simply due to a generalized defect in the responsivity of DIRKO ‐cells to insulin secretagogues, but likely reflects loss of incretin action in the context of GPR119 activation. Moreover, the inability of AR231453 to reduce glucose excursion during an IPGTT in WT mice is not explained by an inability of murine ‐cells to directly respond to this GPR119 agonist.

GPR119 controls gut motility through mechanisms independent of the GLP‐1R, GLP‐2R or Y2R As GLP‐1 regulates glucose homeostasis through control of gut motility [472, 473], and GPR119 activation robustly increases circulating levels of GLP‐1 (Figure 2.3), we examined gastric emptying following administration of AR231453 in WT mice. AR231453 significantly reduced gastric emptying in WT mice (Figure 2.6A). To determine whether the effects of AR231453 on gastric emptying were mediated through a GLP‐1R‐dependent mechanism, we repeated these studies in Glp1r‐/‐ mice. Surprisingly, GPR119 activation continued to reduce gastric emptying in both Glp1r‐/‐ and in DIRKO mice (Figure 2.6A), suggesting that AR231453 controls gastric emptying through a mechanism that does not require the GLP‐1 or GIP receptors. Previous studies of AR231453 demonstrated that this molecule is a highly selective agonist that required the GPR119 receptor for control of glucose tolerance and insulin secretion [377]. To assess whether the inhibitory actions of AR231453 on gastric emptying were also mediated through GPR119, or reflected GPR119‐independent mechanism(s) of action, we examined gastric emptying in GPR119‐/‐ mice. The GLP‐1R agonist exendin‐4 potently reduced gastric emptying in both GPR119+/+ and GPR119‐/‐ mice (Figure 2.6B). In contrast, AR231453 reduced gastric emptying in GPR119+/+ but not in GPR119‐/‐ mice (Figure 2.6B). Hence, GPR119 transduces the actions of AR231453 on gastric emptying.

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Measure plasma acetaminophen

-30 0 15 30 Time (min) -30

Oral AR231453 or vehicle Oral Acetaminophen/ Tail Tail (panels A-C) or IP glucose bleed bleed exendin 4 (panel B) (50ul) (50ul)

Vehicle AR231453 Exendin-4 (n=4) A Vehicle B ** AR231453 7 7 ** * * 6 * 6 * 5 5 4 4 3 3 2 2 AUC (mMAUC /min) AUC (mM min) / 1 1 Plasma Acetaminophen Plasma Plasma Acetaminophen 0 0 WT Glp1r-/- DIRKO Gpr119 +/+ Gpr119 -/- (n=7-8) (n=12) (n=9-10) (n=18) (n=14)

C Vehicle AR231453

5.5 ** p=0.0580 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 AUC (mMAUC / min) 1.0

Plasma Acetaminophen Plasma 0.5 0.0 Glp2r+/+ Glp2r-/- (n=5) (n=6)

Figure 2.6 Gastric emptying in WT, Glp1r‐/‐, DIRKO, Gpr119‐/‐ and Glp2r ‐/‐ mice. Age matched littermate WT and knockout male mice were fasted overnight and given a single dose of either AR231453 (20 mg/kg) , vehicle (A‐C), or exendin‐4 (1ug) (B) before oral administration of a solution of glucose 15% and acetaminophen 1% at a dose of 1.5 g/kg glucose – 0.1 g/kg acetaminophen. Gastric emptying rate was determined as described in methods. Area under the curve for plasma acetaminophen levels in A) WT, Glp1r‐/‐ and DIRKO mice; (B) GPR119+/+ and GPR119‐/‐ mice; (C) Glp2r+/+ and Glp2r‐/‐ mice. Statistical analysis was assessed by ANOVA and paired t‐test, (as appropriate) * p<0.05, ** p<0.01.

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As AR231453 likely increases plasma levels of the related L cell‐derived peptide GLP‐2, and GLP‐2 R activation may also inhibit the rate of gastric emptying [474], we assessed gastric emptying in Glp2r+/+ and Glp2r‐/‐ mice. AR231452 reduced gastric emptying in both Glp2r+/+ and Glp2r‐/‐ mice (Figure 2.6C), demonstrating that the GLP‐2R is not the sole dominant mechanism transducing the AR231453‐dependent inhibition of gastric emptying (Figure 2.6C). GPR119 activation also stimulates local release of PYY from isolated gut mucosa [451], and PYY also inhibits gastric emptying [452]; hence, we examined the effect of acute GPR119 activation on plasma PYY. AR231453 administration prior to oral glucose loading significantly increased plasma PYY in WT mice (Figure 2.7A). Accordingly, we assessed whether the actions of AR231453 on gastric emptying were diminished by concomitant administration of the Y(2) receptor antagonist, BIIE0246, a reagent previously shown to abrogate the effects of PYY on gut motility [452]. AR231453 significantly inhibited gastric emptying despite concomitant administration of BIIE0246 (Figure 2.7B), suggesting that the GPR119‐mediated control of gut motility is not mediated through the Y(2) receptor.

2.5 DISCUSSION

Studies of the biological activity of GPR119 using synthetic receptor agonists initially focused on activity of this receptor in islet cells, demonstrating potent stimulation of insulin release from ‐cells in vitro, and in rodents in vivo [377]. The complexity of GPR119 biology was further expanded to encompass gut peptide action, as pharmacological GPR119 activation also increased levels of both GIP and GLP‐1 in mice and rats [340, 341, 381, 387]. Furthermore, endogenous GPR119 appears essential for maximal GLP‐1 secretion as GPR119‐/‐ mice exhibited reduced levels of circulating GLP‐1 following nutrient or glucose ingestion [387]. These findings raised the possibility that one or both incretin peptides contribute to the glucose lowering actions of GPR119 in the postprandial state. We have now examined the relative importance of three distinct components of GPR119 action that might contribute to the glucoregulatory actions of GPR119 agonists, namely i) incretin peptides, ii) a direct insulinotropic role for GPR119 in islets, and iii) the control of gastric emptying.

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A Tail bleed

- 30 0 5

3 *

2

1 PYY (ng/ml)

0 Vehicle (n=8) AR231453 (n=7)

B

Measure plasma acetaminophen

-45 -30 0 15 45

BIIE0246 or AR231453or Oral Tail Tail V1 (ip) V2 (oral) Acetaminophen/ bleed bleed glucose

V1 - V2 BIIE0246 - V2 V1 - AR231453 BIIE0246 - AR231453

15 * *

10

5 AUC (mMAUC / min) Plasma Acetaminophen Plasma 0 n=7

Figure 2.7 GPR119 activation increases plasma PYY but inhibits gastric emptying independent of the Y2R receptor. Age‐matched male mice were fasted overnight and oral AR231453 (20 mg/kg) or vehicle was administered 30 min before an oral glucose load (OGTT). Blood samples were collected 5 minutes after glucose for the measurement PYY plasma levels (A). For gastric emptying (B) 10 week old C57Bl6 male mice were fasted overnight. At time zero a single ip dose of either vehicle (V1 = 1.2 % DMSO in water) or BIIE0246 (2mg/kg) was administered 15 min before a single oral dose of either vehicle (V2=(80% PEG400, 10% Tween 80, 10% ethanol) or AR231453 (20mg/Kg). A solution of glucose (15%)‐acetaminophen (1%) was administered orally 30 min later at a dose of 1.5 g/kg glucose – 0.1 g/kg acetaminophen and blood samples were collected at 15 and 45 min for assessment of plasma acetaminophen levels as described in methods. Rate of appearance of acetaminophen in plasma as determined by the area under the curve (AUC) (B). Statistical analysis was assessed by t‐test and ANOVA, * p<0.05.

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Our data are consistent with a model invoking all three mechanisms as contributing to the totality of GPR119 action in vivo. We demonstrate that, although the glucoregulatory actions of AR231453 are diminished in DIRKO mice, AR231453 continues to significantly increase plasma insulin levels concomitant with oral glucose loading in DIRKO mice. Hence, an intact incretin axis is not required for the insulinotropic actions of GPR119 agonists in vivo. The failure of AR231453 to lower glycemic excursion in DIRKO mice despite increasing insulin levels may be explained in part by the lack of GIP and GLP‐1 action together with increased plasma levels of glucagon observed in the same experiments. DIRKO mice exhibit modest defects in β‐cell proliferation, and impaired up‐regulation of insulin gene expression and insulin secretion in response to high fat feeding [465]; hence, one explanation for the relatively reduced insulinotropic response to AR231453 in DIRKO mice invokes a potential defect rendering DIRKO islets unable to respond adequately to GPR119 or other insulin secretagogues [465]. Several lines of evidence argue against this interpretation of the data. First, GPR119 expression was comparable in RNA isolated from WT, single incretin receptor knockout, and DIRKO islets. Furthermore, arginine briskly increased plasma insulin levels in both WT and DIRKO mice in vivo. Moreover, DIRKO islets exhibited a significant insulin secretory response to a) AR231453, and b) PACAP in vitro. Hence the available data clearly demonstrate that GPR119 activation likely promotes insulin secretion via two complementary mechanisms; a direct ‐cell effect and by activation of the incretin axis. Interpretation of data from incretin receptor knockout mice must be also tempered by the realization that these mice exhibit compensatory adaptations in levels of circulating incretins and in incretin responsivity [475, 476]. Indeed, we observed a more robust induction of plasma GIP levels with AR231453 in the absence of a functional GLP‐1R, and increased plasma levels of GLP‐1 in Glp1r‐/‐ and DIRKO mice. These findings are consistent with previous studies suggesting that enteroendocrine cells regulate peptide hormone secretion in response to ambient levels of circulating gut hormones [477]. Nevertheless, as previous [478] and more recent [41] studies have demonstrated that classical antagonists used to disrupt GLP‐1 action such as exendin(9‐39) may not be completely specific for the GLP‐1 receptor, the use of mice with genetic disruption of incretin receptor genes provides a valuable complementary model

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for assessing the importance of incretin receptor signaling for glucoregulatory mechanisms activated by engagement of the GPR119 receptor. Previous studies of GPR119 action have not addressed whether glucagon contributes to the glucoregulatory properties observed with GPR119 agonists. GPR119 has been predominantly localized to ‐cells, and in one report to PP cells [479], but not ‐cells. Although AR231453 had no effect on levels of plasma glucagon in WT mice, circulating glucagon levels were surprisingly increased following AR231453 administration in single and double incretin receptor knockout mice. Basal GLP‐1R signaling is known to tonically inhibit glucagon secretion, and even transient blockade of the GLP‐1R using the antagonist exendin(9‐39) increases levels of plasma glucagon following oral glucose challenge in normal and diabetic subjects [480, 481]. Hence, it is possible that loss of the suppressive component of GLP‐1 action on α‐cells in Glp1r‐/‐ and DIRKO mice unmasks a previously undetected action of GPR119, directly or indirectly, on the α‐cell. Nevertheless, it remains unclear how loss of the Gipr also leads to enhanced glucagon secretion following GPR119 activation in the Gipr ‐/‐ mouse. Hence, further exploration of the connection between GPR119 signal transduction and mechanisms regulating α‐cell secretion appears warranted. Considerable previous studies supports the hypothesis that oleoylethanolamide (OEA), an endogenous fatty acid derivative, functions as an endogenous agonist at the GPR119 receptor [337, 341]. Intriguingly, GPR119 expression has been detected in the central nervous system, and OEA reduces food intake [337], raising the possibility that GPR119 functions as a component of a satiety circuit controlling body weight. Similarly, a small molecule GPR119 agonist, PSN632408, also suppressed food intake and reduced body weight in rodents [337]. More recent findings illustrate that the satiety effects of OEA are complex and require CD36 and PPAR [402]. Moreover, OEA retains its anorectic actions in GPR119‐/‐ mice [387]. In contrast, we found no effect of GPR119 activation on food intake using a selective GPR119 agonist in WT, single incretin receptor knockout, or DIRKO mice (Figure 2.8). These findings are consistent with the normal body weight previously reported for GPR119‐/‐ mice [377, 387] and suggest that pharmacological activation or genetic disruption of GPR119 does not produce a phenotype linked to disordered control of energy homeostasis.

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As GPR119 activation significantly increases proglucagon‐derived peptide (PGDP) secretion from the gut L cell and GLP‐1, oxyntomodulin and GLP‐2 all inhibit gastric emptying [461], we hypothesized that enhanced L cell secretion might be associated with reduced gastric emptying. Surprisingly however, there is little previous data examining whether selective GPR119 activation controls gastric emptying. We found that AR231453 significantly inhibited gastric emptying in WT mice. Unexpectedly AR231453 also inhibited gastric emptying in Glp1r‐/‐, DIRKO, and Glp2r‐/‐ mice, demonstrating that the inhibitory effect of GPR119 agonists on gut motility is not strictly dependent on the GLP‐1, GIP or GLP‐2 receptors. Nevertheless, the ability of AR231453 to reduce gastric emptying was lost in GPR119‐/‐ mice, emphasizing that the inhibition of gastric emptying does not reflect an "off target" mechanisms of action of AR231453 and clearly requires a functional GPR119 signaling system. Consistent with our findings, the GPR119 ligand OEA also inhibits gastric emptying in mice, through poorly understood mechanisms independent of PPAR or cannabinoid receptors [450].

Moreover, consistent with our findings using AR231453 in Glp1r‐/‐ mice, the effects of OEA on gut motility were not diminished by co‐administration of the GLP‐1R antagonist exendin(9‐ 39) [450]. More recent experimentation has delineated a role for PYY as a downstream target of GPR119 in the gastrointestinal mucosa. GPR119 activation using the agonist PSN632408 inhibited epithelial electrolyte secretion in a PYY‐ and Y1 receptor‐dependent manner [451]. Unexpectedly, the glucoregulatory and insulinotropic actions of PSN632408 were also attenuated in Pyy‐/‐ mice. Our data extend these findings by demonstrating that AR231453increased plasma levels of PYY in association with enteral glucose loading in vivo. Nevertheless, although exogenous PYY is known to inhibit gastric emptying [482], the inhibitory effects of AR231453 on gastric emptying were not diminished by co‐administration of the Y2 receptor antagonist BIIE0246. Since multiple gut peptides inhibit gastric emptying, and many of these (GLP‐1, GLP‐2, oxyntomodulin, PYY etc) are downstream targets of GPR119, simply inhibiting the action of a single peptide is unlikely to completely reverse the inhibition of gastric emptying observed with GPR119 agonists. Hence, the precise mediators and mechanisms coupling GPR119 activation to control of gut motility require further study.

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A

B

Figure 2.8 The GPR119 agonist MBX3152 (Metabolex) failed to reduce food intake in mice (A) Age matched GLP1r ‐/‐ and littermate controls were kept on RC. Mice were fasted O/N and the day of the experiment mice were given a single oral dose of 30 mg/kg of MBX152 or vehicle. Mice were allowed access to pre‐measured food 30 min after GPR119 agonist dose. Food intake was assessed at 2 and 24 hrs. (B) GLP1r ‐/‐ and +/+ mice on regular chow treated with exendin‐4 were used as positive control.

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In summary, our data illustrate that GPR119 controls enteral glucose tolerance through at least three distinct yet complementary mechanisms. First, activation of the ‐cell GPR119 directly enhances insulin secretion in isolated murine islets independent of incretin receptors. Moreover, we identify a role for GPR119 in the control of gastric emptying that is not completely dependent on the GLP‐1R, GLP‐2R or Y2R. Finally, we demonstrate that functional incretin receptors are required for maximal improvement of oral glucose tolerance following GPR119 activation. Taken together it seems likely that multiple enteroendocrine cell‐derived peptides simultaneously contribute to the glucoregulatory signals and control of gastric emptying emanating from GPR119 activation in the gut. These findings may have implications for optimization of therapeutic strategies and the safety of employing GPR119 agonists for the treatment of type 2 diabetes. For example, a combination therapy of DPP‐4 inhibitors and GPR119 agonists could result in better glucose control for type 2 diabetic subjects.

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Chapter 3

GPR119‐/‐ MICE ARE MORE SUSCEPTIBLE TO STREPTOZOTOCIN‐ INDUCED APOPTOSIS AND TO LIPOTOXIC‐INDUCED ‐CELL FAILURE

G. B. Flock and D. J. Drucker

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3.1 ABSTRACT Activation of G‐protein coupled receptor 119 (GPR119) stimulates insulin secretion in a glucose‐dependent manner in vitro. In vivo, GPR119 agonists transduce their glucoregulatory actions via complementary mechanisms that involve incretin‐dependent glucose‐stimulated insulin secretion and incretin‐independent reduction of gastric emptying. GPR119, like GLP‐1R and GIP‐R, is a GPCR that couples to Gs and stimulates cAMP production in pancreatic ‐cells and intestinal L‐cells. An important feature of β‐cell GPCRs coupled to cyclic AMP generation is their ability to protect the β‐cell from glucolipotoxicity, external injury and ER stress. Furthermore, GLP‐1R activation leads to induction of insulin biosynthesis, β‐cell proliferation, and enhanced β‐cell survival. GPR119 agonists may theoretically protect the β‐cell directly, or indirectly, through stimulation of GLP‐1 secretion. We examined whether activation of GPR119 protects the ‐cell from streptozotocin‐induced apoptosis in vivo in WT mice. Conversely, we investigated if ablation of GPR119 in mice would result in an increased susceptibility to streptozotocin (STZ) compared to WT controls. In addition, we examined the effect of chronic high fat feeding in GPR119‐/‐ compared to WT control mice. Experimental endpoints included: plasma glucose, insulin, glucagon‐like peptide‐1 (GLP‐1) and glucose‐dependent insulinotropic peptide (GIP), pancreatic insulin content, insulin area and islet size distribution. To assess whether endogenous GPR119 signaling is important for ‐cell biology we compared islet mRNA transcript levels of selected key genes involved in ‐cell differentiation, function, proliferation and survival in GPR119‐/‐ and GPR119+/+ mice fed high fat diet (HFD) versus regular chow diet (RC). Our findings indicate that activation of GPR119 does not protect β‐cells from STZ‐induced cytotoxicity. However, GPR119‐/‐ mice are more susceptible to STZ‐induced β‐cell apoptosis and when on HFD, they develop elevated fasting glucose, postprandial glucose intolerance, and impaired ‐cell function. GPR119‐/‐ mice islets showed reduced levels of Irs2, Glp1r, Vipr1 and glucagon mRNA compared to WT islets under basal conditions. Furthermore, on HFD, GPR119‐ /‐ islets failed to increase expression of key genes that play a role in ‐cell function and survival. In summary, our findings provide new insights into the biological importance of GPR119 signaling for β‐cell function and survival.

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3.2 INTRODUCTION Insulin resistance and the resulting increased insulin demand, precedes the development of hyperglycemia [483]. However, β‐cell dysfunction is essential for the development of diabetes [484‐486]. The normal β‐cell response to a chronic excess of nutrients and obesity‐associated insulin resistance is to increase insulin secretion in order to maintain glucose homeostasis. Expansion of functional β‐cell mass [487, 488] and enhanced β‐cell function [489] are important mechanisms for the β‐cell adaptation to increased insulin demand. Eventually, however, sustained insulin resistance results in β‐cell failure associated with an increase in β‐cell death and the development of type 2 diabetes. β‐cell mass is modulated by a fine balance between β‐ cell growth (replication and neogenesis) and β‐cell death (mainly apoptosis). Nutrients are important stimulants of β‐cell mass re‐modeling. For example, increased circulating levels of glucose and free fatty acids (FFA) have been shown to be important determinants of β‐cell mass [487]. An important feature of β‐cell GPCRs coupled to cyclic AMP generation is their ability to protect the β‐cell from cytotoxicity [463]. GLP‐1, in addition to enhancing glucose‐stimulated insulin secretion (GSIS) from pancreatic islets induces insulin biosynthesis [8], β‐cell proliferation, neogenesis and enhances β‐cell survival [386, 490, 491]. The anti‐apoptotic properties of GLP‐1 were demonstrated in diabetic ZDF rats [118], STZ‐induced diabetic mice [132], db/db mice [119] and isolated human islets [133, 134]. Furthermore, GLP‐1R activation protects the β‐cell from the deleterious effects of cytotoxic induced endoplasmic reticulum (ER) stress [394]. Conversely, GLP‐1R knockout mice (Glp1r‐/‐) are more susceptible to β‐cell injury [396] and display abnormal islet size and topography [492]. Moreover, mRNA and protein levels of key genes known to play a critical role for β‐cell cytoprotection, proliferation and survival are down regulated in murine Glp1r‐/‐ islets compared to Glp1r+/+ islets [396]. Several mechanisms are involved in GLP‐1‐mediated β‐cell cytoprotection, including activation of the cAMP–PKA‐CREB pathway [71, 493], reviewed in [120, 494]. GPR119 activation, like GLP‐1R and GIP‐R, couples to Gαs and stimulates cAMP production in pancreatic β‐cells [377] and enteroendocrine L‐cells [341]. GPR119 agonists act as insulinotropic factors enhancing glucose‐stimulated insulin secretion (GSIS) [337, 377, 387, 400]

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and as incretin secretagogues [340, 341, 344]. Hence, GPR119 activation could play an important role in β‐cell survival directly via signaling in the β‐cell or indirectly via stimulation of GLP‐1 secretion. We have now investigated whether activation of GPR119 protects islet β‐cells from STZ‐ induced apoptosis. Conversely, we assessed whether ablation of GPR119 signaling renders the murine β‐cell more susceptible to the deleterious action of STZ and long term high fat (HF) feeding. Furthermore, we investigated whether GPR119 signaling is important for the compensatory response of the endocrine pancreas to high fat feeding. Finally, we evaluated whether GPR119 ‐/‐ islets exhibit differential expression of key genes important for islet β‐cell development and maturity, glucose sensing, insulin biosynthesis, insulin secretion, β‐cell proliferation and survival compared to GPR119+/+ islets; under basal conditions and after high fat feeding. Our studies provide evidence for the importance of endogenous GPR119 signaling for islet adaptation to a high fat diet and STZ‐induced injury.

3.3 RESEARCH DESIGN AND METHOD 3.3.1 Animal Experiments Animal experiments were carried out according to protocols approved by the Mt. Sinai Hospital and the Toronto Centre for Phenogenomics (TCP) Animal Care Committees. Male, age matched, C57BL/6 mice were purchased from in house TCP colony and were allowed to acclimatize to the housing room for 1 wk before experimentation. GPR119‐/‐ and littermate control male mice were on a C57BL/6 genetic background. GPR119‐/‐ mice were obtained from Arena Pharmaceuticals, San Diego, CA [377]. Mice were housed (3–5 per cage) under specific pathogen free conditions in microisolator cages and maintained on a 12‐h light /dark cycle with free access to food and water unless otherwise noted.

3.3.2 Streptozotocin‐Induced Apoptosis For loss of function studies, 8‐10 week old age‐matched GPR119‐/‐ and littermate controls were fasted for 4 h before STZ (50mg/ kg/day) or vehicle (0.1 mol/L sodium citrate, pH5) injection once daily for 5 consecutive days as described [132]. For gain of function studies, 8‐10 week old C57BL/6 male mice (n=8) were randomized to receive twice daily at 9:00 AM and

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5:00 PM an oral suspension of the GPR119 selective agonist AR881 at a dose of 30mg/kg (Arena Pharmaceuticals) or vehicle (0.5% hypromellose (HPMC). AR881 treatment started 3 days prior to STZ treatment and continued during the STZ protocol described above. At the end of the study and 12 h after the last STZ and AR881 dose, mice were euthanized and the entire pancreas was collected, fixed in formalin for 24 h, embedded in paraffin and serial sections were immunostained for insulin and cleaved caspase‐3. Slides were scanned using the ScanScope CS system (Aperio Technologies, Vista, CA). Images were analyzed with ScanScope software (Aperio Technologies) as follows; insulin area was measured manually drawing a line bordering the contour of every islet present in the section and cleaved caspase‐3 immunopositive ‐cells were manually counted. All islets in two independent sections (100 µm apart) of each pancreas were analyzed for insulin and cleaved caspase‐3. For each mouse, the total number of nuclei within insulin positive cells (‐cells) present in 5 (for vehicle) and 10 islets (for STZ) and the number of cleaved caspase‐3 positive cells within those ‐cells were counted, and the insulin area of those islets was measured as described above. To estimate the ‐cell size in GPR119+/+ versus GPR119‐/‐ mice insulin‐immunopositive area (β‐cell area) was divided by the number of beta cells counted within that insulin‐immunopositive area.

3.3.3 High Fat Diet Studies GPR119‐/‐ male mice and age‐matched littermate controls were randomized at 5 weeks of age to receive either standard chow diet (RC) (18% Kcal from fat, Teklad global rodent diet, Harlan laboratories) or high fat diet (45% kcal from fat, Research Diets Inc.). For long‐term HF diet studies, mice were kept on RC or HFD for a total of 34 weeks. Glucose tolerance tests were done at 26 weeks for OGTT and 27 weeks for IPGTT following initiation of HF feeding. Insulin tolerance test (ITT) was done at 31 weeks following initiation of HF feeding. For short‐term HFD studies, mice were kept on a HFD for a total of 13 weeks and islets were collected for qPCR studies.

3.3.4 Glucose Tolerance Tests Mice were fasted overnight, weighed and fasting blood glucose levels were measured by tail vein sampling using a glucometer (Bayer, Toronto, ON). A single oral or ip dose of glucose (1.5 g/kg body weight) was given at time zero. Blood glucose levels were measured by sampling

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from the tail vein of gently held conscious mice, from 5‐120 min after glucose administration. At the 5‐min time point, a blood sample (150 µl) was collected and immediately mixed with 15 µl of a chilled solution containing 5000 kIU/ml Trasylol (Bayer, Toronto, ON), 32 mM EDTA, and 0.01 mM Diprotin A (Sigma St. Louis, MO) and kept on ice for assessment of total GLP‐1, total GIP and insulin in plasma. Plasma was obtained by centrifugation at 4 C and stored at –80 C until determination of insulin (ultrasensitive mouse insulin ELISA, Alpco Diagnostics, Salem, NH), total GLP‐1 (mouse/rat total GLP‐1 assay kit Mesoscale Discovery, Gaithersburg, MD), total GIP (rat/mouse GIP (total) ELISA kit Millipore, Billerica, MA) and glucagon (Milliplex endocrine assay, Millipore).

3.3.5 Insulin Tolerance Test GPR119‐/‐ and control male mice kept on RC or HFD were fasted for 5 hrs. A single dose of insulin (1.2 U / kg) (Humulin R, Eli Lilly, On. Canada) was administered by ip injection and blood glucose was determined at 0, 15, 30, 60 and 90 minutes.

3.3.6 Food Intake Mice were fasted overnight, weighed and then placed in individual cages containing pre‐ weighed food with free access to water. Food was re‐weighed after 2, 4 and 24 hours and food intake was expressed as grams of food consumed per gram of body weight.

3.3.7 Magnetic Resonance Imaging (MRI) For assessment of fat and lean mass body composition, a mouse whole body magnetic resonance analyzer was used (EchoMRI‐100, Echo Medical Systems) according to the manufacturer's instructions.

3.3.8 Indirect calorimetric and locomotor activity Following 24 weeks on HF or RC diet GPR119+/+ and GPR119‐/‐ mice were placed individually in chambers of an Oxymax system (Columbus Instruments) with free access to

water and food for 24 hr, and readings were taken 5 hr after acclimation. Measures of O2

consumption, CO2 production, respiratory quotient, and physical activity were obtained. O2 consumption was measured at 15 min interval and was normalized to body weight. Locomotor activity was assessed as the total distance traveled, calculated from measurement of beam

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breaks in Opto M3 activity monitors (Columbus instruments). Data were collected and analyzed using Oxymax Windows software version 2.3 following manufacturer’s instructions.

3.3.9 Islet Isolation After the mice were euthanized using CO2, the pancreas was inflated via the pancreatic duct with collagenase type V (0.7 mg/mL in Hank’s balanced salt solution), excised, and digested at 37°C for 10–12 minutes. The resulting digest was washed twice with cold Hank’s balanced salt solution (containing 0.25% wt/vol fat free bovine serum albumin) and islets were separated using a Histopaque density gradient. The interface containing the islets was removed, washed twice with Hank’s balanced salt solution containing 0.25% bovine serum albumin. Isolated islets were further purified by handpicking using a light microscope, and were washed once with PBS prior to lysis and RNA isolation with RNeasy Mini kit (Qiagen, Mississauga, ON, Canada) following manufacturer’s protocol.

3.3.10 Complementary DNA Synthesis and Gene Expression Analysis First‐strand complementary DNA was synthesized from total islet or pancreas RNA following the manufacturer’s protocol (RNeasy; Qiagen, Mississauga, ON, Canada) using the SuperScript III reverse transcriptase synthesis system (Invitrogen, Carlsbad, CA) and random hexamers. Quantitative polymerase chain reaction (qPCR) was performed with the ABI Prism 7900 Sequence Detection System using the TaqMan Gene Expression Assays and TaqMan Universal PCR Master Mix (Applied Biosystems, Foster City, CA). Transcripts were normalized to levels of peptidyl‐propyl isomerase A (cyclophilin).

3.3.11 Pancreatic Insulin Content Pancreatic fragments, isolated from the same anatomical portion of each mouse pancreas, were homogenized in ice‐cold acid‐ethanol solution (0.18 m HCl, 70% ethanol). After overnight incubation at 4 C, homogenates were centrifuged at 1600 g for 15 min. The supernatant was collected and stored at ‐20 oC. The pellet was homogenized once more in acid‐ethanol, incubated for 2 h at 4 oC, centrifuged as described above and supernatant collected and combined to previously prepared supernatant. Insulin was determined in these extracts using

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the rat insulin RIA kit (Linco Research, St. Charles, MO) and normalized to protein content, as assessed by the BCA protein assay kit (Pierce, IL; USA).

3.3.12 Islet Morphometric analysis The entire pancreas from GPR119‐/‐ and GPR119+/+ mice from the long‐term HFD study (described above) was dissected, fixed overnight in formalin and embedded in paraffin. Histological sections were immunostained for insulin using the guinea pig anti‐insulin primary antibody (Invitrogen) as previously described [495]. Slides were scanned as described above. Islets present in each section were counted. ‐cell area (µm2) and total pancreatic area (µm2) was measured drawing a line bordering the contour of every islet present in the slide and of the entire pancreatic section. The pancreatic area occupied by ‐cells (insulin positive area) was normalized by total pancreatic area and expressed as percentile. The total number of islets present in each section was normalized by pancreatic area and expressed as percentile. Note: We did not record pancreas weight, thus we were not able to calculate β‐cell mass.

3.3.13 Statistical analysis All results are expressed as mean ± SD with the exception of AUCs which are expressed as Mean ± SEM. Statistical significance was assessed by ANOVA and, where appropriate, Student’s t test using GraphPad Prism (version 4; GraphPad Software). A P value less than 0.05 was considered statistically significant.

3.4 RESULTS GPR119 activation does not protect ‐cells from STZ‐induced apoptosis Glucagon‐like peptide‐1 receptor signaling protects islet β‐cells from apoptosis [132]; this ability is coupled in part to the cAMP‐PKA pathway. We hypothesized that activation of GPR119 will transduce anti‐apoptotic action in pancreatic β‐cells. To determine whether GPR119 activation will protect ‐cells from STZ‐induced apoptosis we treated C57BL/6 mice with the small molecule, specific GPR119 agonist, AR881 (Arena Pharmaceuticals) prior to and concomitant with administration of STZ, 50mg/kg/day as described in methods. We assessed ‐ cell apoptosis immediately following the last STZ dose. Consistent with previous findings, STZ significantly induced ‐cell apoptosis compared to vehicle‐treated mice. However, AR881‐

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treated mice exhibited similar levels of apoptosis following STZ administration as vehicle‐ treated mice (Figure 3.1A). To investigate the efficacy of AR881 we performed an OGTT in C57BL/6 pre‐treated with a single oral dose of AR881 (30mg/kg) or vehicle. AR881 was active and improved glucose tolerance in WT mice as demonstrated by reduction of the area under the glucose excursion curve (AUC) (Figure 3.1B). Glp1r‐/‐ mice are more susceptible to STZ‐induced ‐cell apoptosis [132, 396]. To determine whether ablation of GPR119 would result in an increase in ‐cell susceptibility to STZ, we treated age‐matched GPR119‐/‐ mice and littermate controls with low dose STZ as described in methods, and we assessed ‐cell apoptosis immediately after the administration of the last STZ dose. STZ induced ‐cell apoptosis in both GPR119+/+ and GPR119‐/‐ mice. However, ‐cells from GPR119‐/‐ mice were more susceptible to STZ‐induced cell death compared to WT controls (Figure 3.1C, D). We analyzed all islets present in two independent sections of the entire pancreas of each mouse (n=10‐11). No significant difference in the total insulin positive area (total ‐cell area) analyzed for both GPR119‐/‐ and GPR119+/+ mice treated with STZ was seen (Figure 3.1E). In this study, I calculated the level of STZ‐induced injury by counting the number of apoptotic ‐cells in each islet and normalizing by the measured insulin immunopositive area of each islet. Hence, a difference in ‐cell size between GPR119‐/‐ and WT control mice could introduce an error in the interpretation of the results. To determine whether the size of ‐cells from GPR119‐/‐ mice differs from WT controls, insulin‐immunopositive area (β‐cell area) was divided by the number of beta cells counted within that insulin‐immunopositive area to obtain an average of the area of a β‐cell as previously described [496]. Our results demonstrate that there are no significant differences in ‐cell size among genotypes (Figure 3.1F); hence, the significant increase in the number of STZ‐induced apoptotic β‐cells seen in GPR119‐/‐ compared to WT control islets cannot be attributed to a greater number of ‐cells per unit of insulin area. Furthermore, STZ treatment induced a similar reduction in ‐cell size for both GPR119+/+ and GPR119‐/‐ mice.

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AB 1.75 350 1.50 300

-4 1.25 V 250 1.00 AR881 *

) x 10 200 2 0.02 STZ -cells / insulin / -cells 150  STZ‐AR881 (mM / min) / (mM

AUC glucose 100

area (um 0.01 50 positive Number cleavedcaspase 3 0.00 0 VAR881

CD ) -4 GPR119+/+ (STZ) GPR119‐/‐ (STZ) GPR119+/+ (V)

) x (10 ) x *** 2 2.0 GPR119‐/‐ (V) 1.5 GPR119+/+ (STZ) 1.0 GPR119‐/‐ (STZ) 0.5 0.050

0.025 Cleaved caspase 3 -cells / insulin area (um area / insulin -cells

 0.000 positive

EF )

5 *** 5 150 ** 4 ) x (10 2 100 3 ) / numberof 2

2 -cells  50 1

0 0 Insulin area (um Insulin Insulin area (um

Figure 3.1 Activation of GPR119 does not protect ‐cells from streptozotocin (STZ)‐induced apoptosis in WT mice; however, ablation of endogenous GPR119 signaling increases ‐cell susceptibility to STZ‐induced apoptosis. (A) Cleaved caspase‐3 positive ‐cells normalized by insulin area from C57BL/6 mice treated with vehicle (V) or the GPR119 specific agonist AR881 (n=8). (B) Area under the glucose excursion curve during OGTT for WT mice treated with AR881 (n=7). (C) Cleaved caspase‐3 positive ‐cells normalized by insulin area for GPR119+/+ and GPR119‐/‐ mice treated with vehicle (V) or STZ as described in methods. (D) Immunoreactive cleaved caspase‐3 ‐cells in a typical islet from GPR119+/+ and GPR119‐/‐ mice treated with STZ. (E) Total analyzed pancreatic insulin area. (F) Average area of a ‐cell. * p<0.05, ** p<0.01, *** p<0.001. n = 10‐11 each group.

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GPR119‐/‐ islets exhibit reduced basal levels of mRNA transcripts for genes important for ‐ cell survival, proliferation and function. Islets from Glp1r‐/‐ mice exhibit reduced basal levels of mRNA transcripts for genes important for ‐cell proliferation and survival [396]. To elucidate mechanisms underlying the differential sensitivity to STZ‐induced β‐cell injury of GPR119‐/‐ mice compared to controls, we analyzed RNA levels in islets isolated from 7 weeks old GPR119‐/‐ and WT littermates. We investigated the mRNA abundance of selected genes important for ‐cell survival, proliferation, glucose sensing and metabolism, insulin biosynthesis and secretion, ER stress and autophagy and genes encoding for islet hormones. mRNA transcripts levels of Irs2, Glp1r, Vpac‐1 and glucagon were significantly reduced in islets from GPR119‐/‐ mice compared to WT controls (Figure 3.2). In contrast, no differences were detected in the levels of mRNAs transcripts encoding NKX2, NeuroD, FOXA2, PDX1, AKT1, CREB1 and IGF1, genes important for ‐cell differentiation, maturity and survival (Table 3.1). Moreover, no differences were detected in levels of mRNA transcripts for insulin, pancreatic polypeptide (PP), amylin, somatostatin (SS) and ghrelin (Table 3.1). Likewise, no significant differences in mRNA transcript levels were identified for the islet receptors Gipr, Grpr, CckAr, Egfr, Igf1r, Igf2r, Gpr40, Gpr41, Gpr43, Gpr120 and Pparα (Table 3.1). Furthermore, no significant differences were detected in the mRNA abundance of Glut2, Gck, Ucp2, Kcnj11, Rimbp32 and Vapb, genes important for the regulation of insulin secretion (Table 3.1). We found no differences in the level of mRNA transcripts for gadd153, Bip and Atf4, genes involved in the unfolded protein response (UPR) activated during ER stress. Autophagy is a catabolic process, tightly regulated, by which a starving cell reallocates nutrients from unnecessary processes to more essential processes. Thus, autophagy plays a role in normal cell growth, development and homeostasis. Ubiquitin‐binding protein P62 and autophagy‐related 7 (Atg7) are key regulators of autophagy. We found no differences in the levels of P62 and Atg7 between GPR119+/+ and GPR119‐/‐ mice under basal conditions (Table 3.1).

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Genes with reduced mRNA transcripts levels in GPR119‐/‐ islets compared to WT controls

GPR119+/+ 1 GPR119‐/‐ 1 * * Glp1r IRS-2

0 0

1 1 * * VPAC-1 Glucagon

0 0

Figure 3.2 Levels of mRNA transcripts in islets isolated from GPR119‐/‐ and GPR119+/+ mice. Islets from GPR119‐/‐ mice exhibit reduced levels of IRS‐2, GLP‐1r, Vpac‐1 and glucagon mRNA transcripts compared to WT control mice. mRNA abundance is expressed relative to control mice. GPR119+/+ n=3, GPR119‐/‐ n=4. *p<0.05.

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Table 3.1 Genes expressed at similar levels in GPR119‐/‐ islets compared to WT controls. mRNA abundance is expressed relative to control mice. GPR119+/+ n=3, GPR119‐/‐ n=4.

GPR119+/+ GPR119‐/‐ Gene name Symbol Mean SD Mean SD Activating transcription factor 4 Atf4 1 0.263 1.073 0.317 ATP‐sensitive inward rectifier potassium channel 11 Kcnj11 1 0.326 0.837 0.346 Autophagy‐related 7 Atg7 1 0.162 1.050 0.176 cAMP responsive element binding protein 1 Creb1 1 0.067 0.945 0.260 Cholecystokinin A receptor CckrAr 1 0.578 3.003 2.540 DNA‐damage inducible transcript 3 Gadd153 1 0.344 0.933 0.179 Forkhead box A2 Foxa2 1 0.227 1.280 0.325 Free fatty acid receptor 1 GPR40 1 0.021 0.997 0.137 Free fatty acid receptor 2 GPR43 1 0.265 1.040 0.346 Free fatty acid receptor 3 GPR41 1 0.236 1.082 0.239 Gastrin releasing peptide receptor Grpr 1 0.586 2.706 2.047 Ghrelin Ghrl 1 0.302 3.087 1.805 Glucokinase Gck 1 0.216 0.647 0.104 Glucose transporter 2Glut 2 1 0.250 0.888 0.083 Glucose‐dependent insulinotropic polypeptide receptor Gipr 1 0.174 0.760 0.145 Homeobox protein Nkx‐2 Nkx2 1 0.147 1.347 0.385 Insulin Ins 1 0.185 0.823 0.242 Insulin‐like growth factor 1 Igf1 1 0.821 4.618 4.286 Insulin‐like growth factor 2 receptor Igf2r 1 0.17 1.026 0.076 Insulin‐like growth factor I receptor Igf1r 1 0.127 0.972 0.102 Neurogenic differentiation 1 Neurod1 1 0.424 0.909 0.16 Omega‐3 fatty acid receptor 1 GPR120 1 0.279 1.065 0.31 pancreatic and duodenal homeobox 1 Pdx1 1 0.268 0.985 0.508 Pancreatic polypeptide PP 1 0.595 1.697 0.477 Peroxisome proliferator activated receptor alpha Ppara 1 0.889 1.388 0.813 Prohormone convertase 1/3 Pcsk1 1 0.889 1.083 0.254 Protein kinase B Akt1 1 0.091 0.89 0.145 RIMS binding protein 2 Rimbp2 1 0.371 0.838 0.131 Sequestosome 1 p62 1 0.139 0.923 0.108 Somatostatin Sst 1 0.15 0.717 0.216 Uncoupling protein 2 Ucp2 1 0.138 0.962 0.265 Vesicle‐associated membrane protein, associated protein B and C Vapb 1 0.07 1.089 0.168

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Impaired ‐cell function in high fat diet‐fed GPR119‐/‐ mice Glp1r‐/‐, Gipr‐/‐ and DIRKO mice become glucose intolerant and exhibit defective ‐cell function following HFD feeding [465]. We examined the importance of endogenous GPR119 signaling for the control of glucose homeostasis by placing age‐matched male GPR119‐/‐ and littermate control mice on RC or HFD for 34 weeks beginning at 5 weeks of age. Both GPR119‐ /‐ and WT controls gained significantly more weight when fed HFD compared to RC‐fed mice. Furthermore, GPR119‐/‐ mice on HFD were significantly heavier than WT mice (Figure 3.3A). To interpret these finding we measured food intake and we assessed body composition (fat vs lean mass). Our results indicate that GPR119‐/‐ mice eat less than littermate control mice, however, they exhibited a significant increase in fat mass after prolonged HF feeding (Figure 3.3B, C). The increase in fat mass seen in GPR119‐/‐ following HF feeding was not associated with decreased locomotor activity nor with reduced energy expenditure (Figure 3.3D). To evaluate if prolonged high fat (HF) feeding impairs glucose control in GPR119‐/‐ mice we performed an OGTT after 26 weeks of high fat feeding. GPR119‐/‐ mice exhibited elevated fasting glucose levels (Figure3.3A) and marked glucose intolerance compared to WT mice following HF feeding (Figure 3.3F). Plasma levels of glucose‐stimulated insulin (GSI) were significantly higher for RC‐fed GPR119‐/‐ mice compared to RC‐fed WT mice (Figure 3.3G). However, no further increase in GSI was seen in WT and GPR119‐/‐ mice following HF feeding (Figure 3.3G). The insulin/glucose ratio was increased in HF‐fed compared to RC‐fed WT mice (Figure 3.3H) but not in GPR119‐/‐ mice (Figure 3.3H). No significant difference in pancreatic levels of glucagon were observed in GPR119‐/‐ mice compared to WT controls under both diet conditions (Figure 3.3I). GPR119 agonists require intact incretin receptor signaling for optimal improvement of glucose tolerance in vivo [344]. We examined the circulating levels of glucose‐stimulated GLP‐1 and GIP in RC and HF‐fed GPR119‐/‐ and WT mice. GPR119‐/‐ mice fed RC diet exhibit significantly lower levels of glucose‐stimulated GLP‐1 and reduced, although not significant, plasma GIP levels compared to WT controls (Figure 3.3J). High fat feeding increased glucose

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ABC

** 0.3 1.0 45 *** * 40 WT-RC * 0.8 * 35 KO-RC 0.2 30 WT-HFD 0.6 25 KO-HFD g/g 20 0.4

BW (g) BW * 15 0.1 * 10 * 0.2 5 Food intake / BW (g / g) / (g BW / intake Food 0 0.0 0.0 2 hrs 4 hrs 24 hrs Fat mass / BW Lean/BW D Locomotion Energy expenditure 4500 21000 night 20000 night 4000 WT-RC n=6 3500 19000 WT-HFD n=4 18000 3000 KO-RC n=6 17000 2500 KO-HFD n=6 16000 2000 15000 1500 14000 (kcal/kg/hr) 13000 1000 12000 Ambulatory activity Ambulatory 500 11000 (X + Y beam breaks/hr) 0 10000 7

19 31 7 19 31 Time (hr) Time (hr)

E F 25 7 ** 6 20 * 5 15 4 500 10 3 GPR119+/+ (RC) GPR119+/+ (HFD)

2 (mM) Glucose 5 250

Glucose (mM) Glucose GPR119-/- (RC) 1 GPR119-/- (HFD) 0 0 0 15 30 45 60 75 90 105 120 min) (mM x AUC 0 Time (min)

G H I 0.15 35 1.50 30 1.25 25 * 0.10 1.00 20 0.75 15 0.50 0.05 10 (ng/ml) / (mM) Glucagon (pM) Glucagon Insulin (ng/ml) 0.25 5 Insulin / Glucose 0.00 0.00 0

J 50 ** * * 700 40 600 500 GPR119+/+ (RC) 30 400 GPR119+/+ (HFD) 20 300 GPR119-/- (RC) 200

10 tGIP (pg/ml) GPR119-/- (HFD)

tGLP-1 (pg/ml) 100 0 0

Figure 3.3 GPR119‐/‐ mice on a HFD develop fasting hyperglycemia and fail to control glucose homeostasis during OGTT. Age‐matched GPR119+/+ (WT) and GPR119‐/‐ (KO) male mice were kept on either RC or HFD for 26 weeks prior to OGTT. (A) Body weights at the time of OGTT, (26 weeks on HFD). (B) Food intake, (6 months on HFD). (C) Body composition assessed by MRI (5 months on HFD). (D) Locomotor activity and energy expenditure (oxygen consumption, VO2) assessed by indirect calorimetric using an Oxymax system (5 months on HFD). (E) Glucose levels following 15hr fasting. (F) Glucose excursion during OGTT; inset area under the curve (AUC). (G) GSI. (H) GSI normalized by circulating glucose levels during OGTT. (I) Plasma glucagon levels. (J) Glucose‐stimulated plasma incretins. (n=10‐11) *p<0.05, **p<0.01, ***p<0.001. GPR119+/+ (RC) n=9, GPR119+/+ (HFD) n=12, GPR119‐/‐ (RC) n=6, GPR110‐/‐ (HFD) n=9.

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stimulated GLP‐1 and GIP levels in GPR119‐/‐ mice but not in GPR119+/+ mice compared to RC‐ fed mice (Figure 3.3J). To assess ‐cell function independent of the entero‐insular axis, we performed an IPGTT in HF and RC‐fed GPR119‐/‐ and littermate‐control mice. GPR119‐/‐ mice fed a HFD were glucose intolerant during IPGTT (Figure 3.4A). Furthermore, insulin levels relative to circulating glucose levels were significantly lower in HF‐fed GPR119‐/‐ mice compared to RC‐fed GPR119‐/‐ mice, (Figure 3.4B) despite a significant increase in the area under the glucose curve (Figure 3.4A, inset). Hence, GPR119‐/‐ mice, following HF feeding, become glucose intolerant and failed to

appropriately increase insulin levels during oral and IP glucose challenge. Glucagon levels following IP glucose administrations were similar among genotypes under both diet conditions (Figure 3.4C). We next examined insulin actions by performing an insulin tolerance test (ITT) in GPR119‐/‐ and WT controls after 31 weeks of HF or RC feeding. The levels of glucose following 5 h fasting and before insulin administration were significantly higher in GPR119‐/‐ mice on HFD compared to all other groups (Figure 3.5A, inset). To correct for this difference in Figure 3.5B we expressed glucose levels during ITT relative to fasting glucose as a percentile. No difference in the sensitivity to exogenous insulin was seen among all groups;

(Figure 3.5B). We examined whether the islet compensatory response to HF feeding was compromised in GPR119‐/‐ mice. In this study, WT and GPR19‐/‐ mice did not display a significant increase in the relative β‐cell area (Figure 3.6A) and in the number of islets (Figure 3.6B), following 34 weeks of HFD compared to RC‐fed mice. We then examined whether endogenous GPR119 signaling is required for maintenance of normal islet topography under RC and HFD. Analysis of the islet size distribution reveals that both GPR119‐/‐ and WT mice increased the percentage of large islets (> 20000 um2) when fed a HFD (Figure 3.6C). Conversely, a significant reduction in the percentage of small islets is seen for both GPR119‐/‐ and WT mice following prolonged HF feeding (Figure 3.6C). Furthermore, the overall relative contribution of islets by size (um2) to the total insulin area demonstrates an increase in large islets (> 20000 m2) and a decrease in single cell size islets (< 300 m2) for GPR119‐/‐ and WT mice on HFD (Figure 3.6D). Hence, it appears that endogenous GPR119 signaling is not essential for control of islet size topography.

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* 15 ** A ) 10 2

30 x (10 5 AUC (mM x min) 0

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10 GPR119+/+ (RC) Glucose (mM)Glucose GPR119+/+ (HFD) GPR119-/- (RC) GPR119-/- (HFD) 0 0 15 30 45 60 75 90 105 120 Time (min)

B GPR119+/+ (RC) 0.25 GPR119+/+ (HFD) * GPR119-/- (RC)

GPR119-/- (HFD) 0.20

0.15

0.10 (ng/ml) / (mM) / (ng/ml) Insulin / Glucose 0.05

0.00

C 35 30 25 20 15 10 Glucagon (pM) 5 0

Figure 3.4 GPR119‐/‐ mice fed a HFD exhibit impaired ‐cell function. (A)Glucose excursion during IPGTT and area under the curve (AUC), inset. (B) Glucose‐stimulated insulin relative to glucose levels during IPGTT. (C) Glucagon levels. (RC) n=9, GPR119+/+ (HFD) n=12, GPR119‐/‐ (RC) n=6, GPR110‐/‐ (HFD) n=9 *p<0.05, **p<0.01, ***p<0.001.

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A 17.5 15.0 GPR119+/+ (RC) 12.5 GPR119+/+ (HFD) GPR119-/- (RC) 10.0 GPR119-/- (HFD) 7.5

Glucose (mM) 5.0 2.5 0.0 0 25 50 75 100 125 150 Time (min)

B

120

90

60

30 Relative Glucose (%) Relative Glucose

0 0 25 50 75 100 125 150 Time (min)

8000 GPR119+/+ (RC) GPR119+/+ (HFD) 6000 GPR119-/- (RC) GPR119-/- (HFD) 4000 AUC (% x min) 2000 % fasting glucose fasting % 0

Figure 3.5 GPR119‐/‐ and WT control mice, chronically fed HFD do not display greater insulin resistance as measured by the insulin tolerance test (ITT). (A) Plasma glucose. (B) Plasma glucose levels during ITT expressed relative to fasting glucose (percentage). (RC) n=9, GPR119+/+ (HFD) n=12, GPR119‐/‐ (RC) n=6, GPR110‐/‐ (HFD) n=9.

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AB

1.4 GPR119+/+ (RC) ) -6 2.0 1.2 GPR119+/+ (HFD) GPR119-/- (RC) 1.0 1.5 GPR119-/- (HFD) 0.8 1.0 -cell area(%)

 0.6

0.4 0.5 0.2 Relative 0.0

0.0 numberRelative islet ( 10 x

C D ) 2 60 60 * +++ 50 50 40 + 30 40 20

-cell area (um 10 30  2.0 (%) ) /

+ 2 1.5 Islets (%) 20 * + 1.0 + 10 * * 0.5 0 0.0 ll e 2 le ll Islet size (um le Islet area (um ) a m ge g a um rg iu r n m i ing d i s la la s s 0 sm 0 0 00 0 me 00 0 med 00 00 -5 0 3 00 0 0 00 < 0 < 3 0 0 >2 30 >20 300-5 2 0- 0 E 50 F 5000-20 3 0.8 0.7

2 g) 0.6  0.5 0.4 1 0.3 Relative Ins2 Relative

content(ng/ 0.2 0.1 Relative pancreatic insulin pancreatic Relative 0 0.0

Figure 3.6 No significant difference in islet size topography, number of islets, levels of insulin transcripts and pancreatic insulin content in GPR119‐/‐ compared to GPR119+/+ mice. (A) Insulin‐immunopositive area relative to pancreatic area, (percentage). (B) Number of islets expressed relative to pancreatic area (m2). (C) Islet size distribution. (D) Contribution of islet area (by islet size) to total insulin‐ immunopositive area (percentage) (E) Insulin mRNA level expressed relative to GPR119+/+ (RC) value. (F) Pancreatic insulin content (ng) normalized by total protein (µg). RC (regular chow diet), HF (high fat diet).

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We investigated whether GPR119‐/‐ mice exhibit impaired up‐regulation of insulin gene expression and reduced pancreatic insulin content following 34 weeks of HF feeding compared to WT control mice. Levels of pancreatic insulin mRNA transcripts were not increased in HF‐fed WT and GPR119‐/‐ mice compared with those of RC‐fed mice (Figure 3.6E). An increase, although not significant, in pancreatic insulin content was seen in WT and GPR119‐/‐ mice following HF feeding compared to mice fed RC (Figure 3.6F). However, no significant difference in the levels of insulin mRNA and pancreatic insulin content, under both feeding conditions, was observed among genotypes (Figure 3.6E, F). Furthermore, no significant differences in mRNA transcripts levels for glucagon, pancreatic polypeptide (PP) and somatostatin were seen in the pancreas of GPR119‐/‐ vs WT mice following HF or RC feeding (Figure 3.7). Interestingly, HFD induced significant up‐regulation of amylin (Iapp) mRNA levels in the pancreas of GPR119‐/‐ mice but not in WT mice (Figure 3.7). To elucidate mechanisms underlying the impaired ‐cell function seen in GPR119‐/‐ mice fed HFD we analyzed the levels of mRNA transcripts in islets isolated from WT and GPR119‐/‐ mice following short term HF feeding. Mice were started on HFD at 5 weeks of age, body weights were measured weekly, and fasting glucose was assessed at 4 and 12 weeks after initiation of HF feeding. HF fed mice gained significantly more weight compared with RC fed mice, however no differences among genotypes were observed (Figure 3.8A). HF‐fed GPR119‐/‐ mice exhibit significantly higher fasting glucose levels compared to RC‐fed mice. Moreover, GPR119‐/‐ mice fed a HFD display moderately higher fasting glucose levels compared to those for HF‐fed WT mice (Figure 3.8B) and they develop impaired ‐cell function as assessed by IPGTT 8 weeks after initiation of HF feeding. Once ‐cell impairment was demonstrated by IPGTT (Figure 3.8C) islets were isolated for qPCR studies following 13 weeks of HFD. We investigated the levels of expression of selected genes important for β‐cell differentiation, maturity, function and survival, ER stress and autophagy. Remarkably, islet mRNA transcript levels for Glp1r, Gipr, CckAr, Igf2r, Igf1, Irs2, AKT1, Gck, amylin, Gpr40, Gpr43 and Gpr120 were significantly increased in WT but not in GPR119‐/‐ mice islets following HF‐feeding (Figure 3.9A). Conversely, HF feeding significantly reduced mRNA levels of Grpr and Kcnj11 in GPR119‐/‐ islets but not in WT controls (Figure 3.9A).

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Relative transcript levels of genes encoding for pancreatic hormones

3.5 4.5 GPR119+/+ (RC) 4.0 3.0 GPR119+/+ (HFD) 3.5 2.5 GPR119-/- (RC) 3.0 GPR119-/- (HFD) 2.0 2.5 PP Gcg 1.5 2.0 1.5 1.0 1.0 0.5 0.5 0.0 0.0

2.0 3.5 * 3.0 1.5 2.5 2.0 1.0 Sst Iapp 1.5

0.5 1.0 0.5 0.0 0.0

Figure 3.7 No significant difference in the transcript levels of glucagon (Gcg), pancreatic polypeptide (PP) and somatostatin (Sst) in pancreas of GPR119‐/‐ compared to GPR119+/+ following HF feeding. Following 34 weeks of HFD, mice were euthanized and pancreas was collected for the assessment of mRNA abundance by qPCR. Gene expression levels for each transcript are expressed relative to GPR119+/+ (RC). *p<0.05.

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A 40

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20 GPR119+/+ (RC) GPR119+/+ (HFD) 10 GPR119-/- (RC) Body weightBody (g) GPR119-/- (HFD) 0 0 1 2 3 4 5 6 7 8 9 10 11 12 13 Weeks B

9 ** GPR119+/+ (RC) 8 GPR119+/+ (HFD) * 7 GPR119-/- (RC) 6 GPR119-/- (HFD) 5 4 3 Glucose (mM)Glucose 2 1 0 4 weeks 12 weeks

C

1000 * 35 750 500 30 AUC 250 (mM x min) 25 0 20 15 10 Glucose (mM)Glucose 5 0 0 10 20 30 40 50 60 70 80 90 100 Time (min)

Figure 3.8 GPR119‐/‐ mice fed a HFD develop fasting hyperglycemia and fail to control glucose homeostasis during IPGTT. (A) Weekly body weights. (B) Fasting plasma glucose following HF feeding. (C) Glucose excursion during IPGTT at 8 after initiation of HFD feeding; inset, area under the glucose curve (AUC). Regular chow diet (RC), high fat diet (HFD). GPR119+/+ (RC) n=3, GPR119+/+ (HFD) n=4, GPR119‐/‐ (RC) n=4, GPR119‐/‐ (HFD) n=5. *p<0.05, **p<0.01, ***p<0.001.

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A mRNA transcripts regulated by HF feeding in WT compare to GPR119‐/‐ islets

*** * *** GPR119+/+ (RC) GPR119+/+ (HFD) Gipr Glp1r CckAr GPR119-/- (RC) GPR119-/- (HFD)

*** *** *** *

1 Irs2 Igf1 Igf2r AKT1

0

*** * *** * *** Gck GPR43 GPR40 Amylin GPR120

* * * * Bip Grpr Kcnj11

B mRNA transcripts downregulated in GPR119‐/‐ islets

*** ** * *** *** CckBr Ghrelin Adcyap1

Figure 3.9 Islet levels of mRNA transcripts in GPR119‐/‐ and +/+ mice fed a HF diet as determined by qPCR. mRNA abundance is expressed relative to RC‐fed GPR119+/+ mice (WT‐RC). (A) mRNA transcripts differentially up‐ regulated by HF D. (B) mRNA transcripts down‐regulated in GPR119‐/‐ mice. GPR119+/+ (RC) n=3, GPR119+/+ (HFD) n=4, GPR119‐/‐ (RC) n=4, GPR119‐/‐ (HFD) n=5. *p<0.05, **p<0.01, ***p<0.001.

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These findings raise the possibility that loss of endogenous GPR119 signaling may render islets unable to respond to the challenge imposed by HF feeding. Interestingly, islets from RC‐ fed GPR119‐/‐ mice exhibit significantly lower expression levels of Cckbr, ghrelin and adenylate cyclase activating polypeptide 1 (Adcyap1) compared to those from RC‐fed WT mice (Figure 3.9B). Furthermore, HF feeding resulted in a significant down‐regulation of mRNA transcript levels of Adcyap1 and ghrelin in islets from WT mice but not from GPR119‐/‐ mice (Figure 3.9B). Fatty acids induce ER stress and β‐cell death [497, 498]. We investigated whether high fat feeding differentially affects levels of mRNA transcripts encoding for proteins involved in ER stress and the unfolded protein response in GPR119‐/‐ mice compared to WT controls by examining the levels of Bip and Gadd153 transcripts. BIP is involved in the folding and assembly of proteins in the endoplasmic reticulum (ER); and plays a role in the transport of aberrant proteins destined for degradation across the ER membrane. Bip synthesis is markedly induced under conditions that lead to the accumulation of unfolded polypeptides in the ER such as high fat feeding. Our findings demonstrate that HF feeding induced to the same extent Bip mRNA levels in GPR119‐/‐ and WT islets and no significant differences were found among genotypes (Figure 3.9A). No differences were observed in the levels of Gadd153 under both diets and among genotypes (Table 3.2). Table 3.2 shows data for mRNA transcripts of selected genes important for β‐cell differentiation, maturity, function and survival, ER stress and autophagy with similar levels among genotypes and with no change following HF feeding.

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Table 3.2 Islet mRNA transcripts with no change following HF feeding mRNA abundance is expressed relative to RC‐fed GPR119+/+ mice (WT‐RC). GPR119+/+ (RC) n=3, GPR119+/+ (HFD) n=4, GPR119‐/‐ (RC) n=4, GPR119‐/‐ (HFD) n=5.

Gene name Symbol WT‐RC WT‐HFD KO‐RC KO‐HFD Mean SD Mean SD Mean SD Mean SD Autophagy‐related 7 Atg7 1 0.040 1.163 0.295 1.129 0.291 0.867 0.116 cAMP responsive element binding protein 1 Creb1 1 0.126 1.061 0.318 1.048 0.191 0.909 0.148 DNA‐damage inducible transcript 3 Gadd153 1 0.064 1.189 0.290 1.349 0.299 1.128 0.203 Forkhead box A2 Foxa2 1 0.254 0.964 0.290 1.002 0.272 0.906 0.189 Free fatty acid receptor 3 GPR41 1 0.128 0.989 0.292 0.929 0.240 0.876 0.185 Glucagon Gcg 1 0.361 0.674 0.164 1.068 0.486 0.529 0.107 Glucose transporter 2Glut 2 1 0.151 0.971 0.322 1.171 0.201 0.875 0.087 G‐protein coupled receptor 119 GPR119 1 0.277 1.608 0.411 0.000 0.000 0.000 0.000 Homeobox protein Nkx‐2 Nkx2 1 0.171 0.859 0.223 0.729 0.119 0.761 0.265 Insulin Ins 1 0.027 1.124 0.444 1.191 0.247 1.257 0.288 Insulin receptor Insr 1 0.072 1.052 0.356 1.007 0.183 0.867 0.108 Neurogenic differentiation 1 Neurod1 1 0.061 1.141 0.291 1.155 0.198 0.942 0.097 pancreatic and duodenal homeobox 1 Pdx1 1 0.152 1.098 0.424 1.099 0.099 1.689 1.085 Pancreatic polypeptide PP 1 0.158 0.529 0.291 0.871 0.477 0.421 0.244 Peroxisome proliferator activated receptor alpha Ppara 1 0.275 1.909 1.718 1.299 0.811 2.667 1.986 Prohormone convertase 1/3 Pcsk1 1 0.098 1.352 0.356 1.097 0.146 1.190 0.228 RIMS binding protein 2 Rimbp2 1 0.082 1.268 0.494 1.184 0.428 0.843 0.213 Sequestosome 1 p62 1 0.081 1.123 0.344 1.040 0.188 1.062 0.142 Somatostatin Sst 1 0.194 1.019 0.334 1.189 0.626 0.807 0.163 Uncoupling protein 2 Ucp2 1 0.142 1.248 0.283 1.182 0.093 1.085 0.106 Vesicle‐associated membrane protein, associated protein B and C Vapb 1 0.043 1.242 0.290 1.173 0.163 1.033 0.108

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3.5 DISCUSSION Type 2 diabetes is characterized by hyperglycemia, insulin resistance and relative insulin deficiency. Moreover, type 2 diabetes is associated with cardiovascular complications, retinopathy and kidney disease and improving glycemic control is critical in reducing the risk of associated complications [499]. However, a major challenge in the treatment of type 2 diabetes is that despite pharmacological interventions, glycemic control deteriorates over time [500]. Insulin resistance is not sufficient for the development of type 2 diabetes. In fact, before the development of hyperglycemia and type 2 diabetes, insulin resistance is counterbalanced by islet adaptation. The islet‐adaptive response involves increased insulin secretion and β‐cell mass [501]. However, a prolonged increased in systemic insulin demand is deleterious to β‐cells leading to β‐cell functional impairment and decrease in islet number and size due to an increase in β‐cell apoptosis [501‐504]. Hence, approaches to protect the β‐cells and to improve β‐cell function are desirable strategies in the treatment and prevention of type 2 diabetes. GPR119 activation leads to increased GSIS and GLP‐1 secretion via an increase in cAMP accumulation [337, 376, 377]. However, the importance of GPR119 signaling for β‐cell cytoprotection and survival has not been determined. Our gain of function studies employing the GPR119‐specific synthetic ligand, AR881, demonstrate that opposite to what has been seen for incretin receptor agonists [132, 396, 461], sustained pharmacological activation of GPR119 fails to protect β‐cells from STZ‐induced apoptosis (Figure 3.1A). It is important to consider aspects of the experimental design. We have used the small molecule AR881 provided by Arena Pharmaceuticals following the manufacturer’s recommended dosage schedule. However, we do not know the AR881 pharmacokinetic profile and no publications using this compound are currently available. We have tested the efficacy of a single dose of AR881 in regulating glucose homeostasis (Figure 3.1B); however, we lack information on the efficacy of sustained administration of AR881. Furthermore, the natural ligand OEA and the specific synthetic ligands PSN632408 and PSN375963 all showed bell‐shape dose response profiles in vitro [337, 341, 505] which indicates the possibility of receptor

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desensitization. Thus, sustained GPR119 activation by AR881 could induce receptor desensitization in vivo, resulting in absence of AR881‐mediated cytoprotective actions. Furthermore, it is possible that our experimental protocol could have resulted in down‐ regulation of GPR119 mRNA transcripts or protein levels. Hence, further studies are necessary to assess if activation of GPR119 plays any role preventing STZ‐induced injury are needed. Our loss of function studies using GPR119‐/‐ mice clearly demonstrates that lack of endogenous GPR119 signaling renders the β‐cell more susceptible to STZ–induced apoptosis (Figure 3.1C, D). Abrogation of endogenous GLP‐1 signaling results in an increased susceptibility to STZ‐induced β‐cell apoptosis [396]. GPR119‐/‐ mice exhibit reduced circulating GLP‐1 levels [387]. Hence, it is possible that the enhanced sensitivity to STZ‐induced injury seen in GPR119‐/‐ β‐cells could be the result of direct abrogation of GPR119 signaling, reduced GLP‐1 action in the β‐cell or a combination of both. Further studies are necessary to elucidate the mechanisms involved. Islets from Glp1r‐/‐ mice exhibit reduced mRNA transcript levels of epidermal growth factor receptor (Egfr), Irs2 and Igf1r [396, 506]. Furthermore, compensatory mechanisms were observed in islets of the global : glucagon‐like peptide‐1 receptor double knockout (Gcgr‐/‐:Glp1r‐/‐) mice and in DIRKO mice treated with Gcgr antisense oligonucleotides (Gcgr Asos). In these mice, islets exhibit increased cholecystokinin A receptor (CckAr) and GPR119 mRNA transcripts. Accordingly, doses of AR231453 and of CCK that failed to improve glucose tolerance in WT, Gcgr‐/‐ and Glp‐1r‐/‐ mice, significantly induced insulin release and improved glucose control during OGTT in Gcgr‐/‐:Glp1r‐/‐ mice [507]. We have shown that islets isolated from young GPR119‐/‐ mice, maintained under basal conditions (RC diet), exhibit similar mRNA transcript levels compared to GPR119+/+ islets for the majority of the molecules investigated (Table 3.1). However, interestingly, we saw reduced levels of mRNA transcripts of Irs2, Glp1r, Vipr1 and Gcg (Figure 3.2A). Insulin receptor substrate‐2 (IRS2) plays a critical role in β‐cell cytoprotection and proliferation [508]. Furthermore, GLP‐1 stimulates insulin mRNA expression, β‐cell proliferation and prevents β‐cell apoptosis via several mechanisms engaging IRS‐2‐mediated signal transduction [509]. In addition, studies in islets isolated from young and old, lean and obese

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(ob/ob) mice suggested that impaired VIP signaling may contribute to the development of impaired glucose‐stimulated insulin secretion during aging and the development of type 2 diabetes [510]. Moreover, transgenic mice with targeted overexpression of human VIP in islet β‐cells exhibit reduced postprandial glycemia and improved glucose tolerance [511]. In contrast, global VIP knockout mice display elevated, both postprandial and fasting, glucose levels compared with WT mice [512]. Hence, our findings indicating that GPR119‐/‐ islets exhibit significantly lower transcript levels for Irs2 and the insulinotropic Glp1r and Vipr1 receptors compared to GPR119+/+ islets may partially explain the increased susceptibility of GPR119‐/‐ β‐cells to STZ and HFD‐induced glucose intolerance in GPR119‐/‐ mice. Interestingly, we identified reduced glucagon mRNA levels in GPR119‐/‐ islets isolated from 7 week old mice compared to WT controls. However, we have not seen differential Gcg expression in islets isolated from 17 week old mice and no difference was observed in Gcg mRNA levels in pancreas isolated from 38 week old GPR119‐/‐ mice compared to WT controls under basal and HF feeding conditions. In situ hybridization studies suggested that a subset of glucagon‐producing cells could express GPR119 in murine pancreas [377] and GPR119 transcripts were identified by qPCR in the TC1‐6 cell line [33]. However, subsequent immunofluorescent studies of rat and mouse pancreatic sections indicated that GPR119 expression co‐localizes primarily to insulin producing cells [377]. Interestingly, we have shown in chapter 2 that in single and double incretin receptor knockout mice, GPR119 activation by AR231453 increased glucagon levels during OGTT. It is currently not known if endogenous GPR119 signaling plays any role in the modulation of α‐cell function and the broader biological significance of our findings is not known. PDX1 is essential for GLP‐1‐mediated β‐cell proliferation, cytoprotection and survival (reviewed in [120]. GLP‐1 transduces these actions via mechanisms that promote PDX1 translocation into the nuclei where it regulates transcription of target genes important for β‐ cell function and survival [120, 136]. We have not observed differences in the mRNA level of PDX1 in GPR119‐/‐ islets compared to WT controls. However, PDX1 nuclear exclusion could result in enhanced susceptibility to β‐cell injury independent of changes in mRNA transcript levels. It is important to mention that the studies presented in this chapter are based on the analysis of mRNA transcript levels; hence, further studies to investigate differences in protein

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expression levels, protein activation and protein intracellular localization between GPR119‐/‐ and GPR119+/+ islets are needed. Consistent with our results using the STZ‐induced β‐cell injury model, GPR119‐/‐ mice were more susceptible to the deleterious effects of HF feeding. GPR119‐/‐ mice fed HFD gained significantly more weight than WT controls (Figure 3.3A). The difference in BW following HFD among genotypes could not be explained by an increase in food intake in GPR119‐/‐ mice compared to WT controls. Furthermore, GPR119‐/‐ mice did not exhibit reduced energy expenditure nor reduced locomotor activity. GLP‐1 has been shown to reduce triglyceride absorption and apolipoprotein production in rats [513]. In addition, DPP‐4 inhibitors were shown to reduce postprandial lipoprotein levels in hamsters, mice and humans, predominantly through inhibition of intestinal chylomicron secretion. In addition, Glp1r‐/‐ mice display increased circulating triacylglycerol‐rich lipoprotein (TRL) following olive oil oral gavage compared to control mice [514]. GPR119‐/‐ mice exhibit reduced glucose‐stimulated GLP‐1 levels; hence, they could exhibit increased intestinal TRL absorption compared to WT mice. Further studies are needed to investigate the mechanisms associated with the increased in body weight in GPR119‐/‐ mice compared to WT control following prolonged HF feeding. High‐fat feeding resulted in an impairment of β‐cell function in GPR119‐/‐ mice compared to littermate controls as demonstrated by an increase in fasting glucose levels and a failure to control glucose excursions associated with insufficient insulin secretion during glucose tolerance tests (GTT) (Figure 3.3E‐H and Figure 3.4A, B). The HFD‐induced impairment in glucose in GPR119‐/‐ mice could not be attributed to differentially elevated glucagon levels following OGTs (Figure 3.3I and Figure 3.4C), nor to differential insulin sensitivity as assessed by ITT (Figure 3.5A, B). However, one limitation of ITT as a method to evaluate insulin resistance is its low precision. Thus, a better method will be performing a euglycaemic insulin clamp. Our results are in contrast with previously published data [387]. Using a different GPR119‐/‐ mouse line than the one used in our lab, Lan et al demonstrated that the weight gain of GPR119‐/‐ mice fed a HFD (45% Kcal from fat) was comparable to WT controls. No differences in fasting glycemia were observed among genotypes. Furthermore, OGTT did not reveal differences due to genotype. Interestingly and in accordance with previously published data [387], our results

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reveal that endogenous GPR119 signaling is important for physiological incretin secretion as GPR119‐/‐ mice exhibit significant reduction of glucose‐stimulated GLP‐1 levels (Figure 3.3J). Although not significant, GIP levels were also lower compared to WT controls under basal conditions (RC diet) (Figure 3.3J). We demonstrated that GPR119‐/‐ mice fed a HFD develop fasting hyperglycemia. It has been shown that GPR119 agonists stimulate GLP‐1 secretion via glucose‐independent mechanisms [515]. Furthermore, there is evidence indicating a role for GLP‐1 in the regulation of circulating fasting glucose and glucagon levels [516, 517]. In our studies, we have not investigated the fasting levels of GLP‐1 and GIP. However, if GPR119‐/‐ mice exhibit reduced tonic incretin release, this could contribute to the HFD‐induced fasting hyperglycemia observed in our studies. Surprisingly, disruption of GPR119 signaling did not affect islet β‐cell adaptation to HF feeding. Indeed, no significant differences in β‐cell area, islet size, and pancreatic insulin mRNA and protein content were observed among genotypes (Figure 3.6A‐F). Furthermore, HF feeding induced a comparable increase in the number of large islets in both GPR119‐/‐ and +/+ mice (Figure 3.6C, D). It is important to mention that in this study, WT mice failed to increase β‐cell area and insulin content in response to HF feeding. One possible explanation could reside in the composition of the RC diet used that contains 18% Kcal from fat sources. Hence, pancreatic islets from WT mice fed RC could have partially adapted to the 18% fat, masking further effects associated with the HFD (45% Kcal from fat). We are currently repeating this study using a RC diet that contains 10% of Kcal from fat vs HFD (45% Kcal from fat). Our analysis of mRNA transcripts levels from GPR119‐/‐ and +/+ pancreas under basal (RC) and HF feeding conditions revealed no genotypic differences and no effect of diet on mRNA for Gcg, PP, and Sst (Figure 3.7). Surprisingly, amylin mRNA levels were significantly induced by HF feeding in the pancreas of GPR119‐/‐ but not WT control mice (Figure 3.7). Our studies are the first to investigate islet gene expression in the context of HF feeding in a model of genetic disruption of endogenous GPR119 signaling. Our results reveal a critical role for GPR119 signaling transducing β‐cell‐adaptive transcriptional responses to HF feeding. HF feeding induced mRNA levels of Glp1r, Gipr, CckAr, Igf2r, Gpr40, Gpr43, Gpr120, Gck, Akt1, Irs2 and Igf1 in WT mice islets. In contrast, no change was observed in islets from HF‐fed GPR119‐/‐ mice

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(Figure 3.9A). As mentioned above, HF feeding induced a massive increase in islet‐CckAr mRNA levels in WT mice compared to mice kept on a RC diet. Interestingly, studies of murine islet adaptation to physiological challenges such as pregnancy‐induced insulin resistance have revealed numerous changes in mRNA transcript levels, among them up‐regulation of GPR119 and CckAr gene expression [518]. Furthermore, studies using DIRKO and Gcgr‐/‐Glp1r‐/‐ double knockout mice revealed compensatory mechanisms involving up‐regulation of CckAr and GPR119 mRNA expression and enhanced sensitivity to CCK and AR231453 treatment [507]. The biological implication of these findings is currently unknown. However, the ob/ob‐CckLacz mice (where the gene encoding for Cck has been replaced by the LacZ gene)[519] exhibit increased β‐cell apoptosis compared to ob/ob‐CckWT mice, as assessed by TUNEL staining of pancreatic sections [519]. In addition, it has been shown that the OLETF rat which carries a null CckAr mutation, develops obesity‐induced diabetes [520]. As with the ob/ob‐CckLacz mice, OLETF rats cannot expand their ‐cell mass due to increased ‐cell death; thus they cannot compensate for peripheral insulin resistance. Hence, it is believed that in rodents the prosurvival effect of CCK is mediated via the CCKAR [519]. Based on our current knowledge we could hypothesize that while WT mice up‐regulate CckAr mRNA levels, thus mounting a compensatory mechanism believed to protect ‐cells from HFD‐induced injury, GPR119 mice failed to do so, rendering β‐ cells more susceptible to HFD‐induced apoptosis. We have detected HFD‐induced down‐regulation of Kcnj11 mRNA transcripts (also known as Kirk6.2) in GPR119‐/‐ but not in WT mice fed HFD. We do not know if the reduction in mRNA levels translates to reduced KATP channel expression and/or function in GPR119‐/‐ islets. However, if this is the case, it could have implications for impairing GSIS in GPR119‐/‐ β‐cells. Taken together our findings provide new information on the importance of basal GPR119 receptor action for β‐cell function and survival. Moreover, our findings imply that elimination of endogenous GPR119 signaling, directly or indirectly, modifies susceptibility to β‐cell injury in association with perturbation in levels of key signaling molecules important for β‐cell function and survival.

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Chapter 4

PROGESTERONE STIMULATES GLP‐1 SECRETION VIA MEMBRANE PROGESTERONE RECEPTORS YET IMPROVES GLUCOSE TOLERANCE VIA A GLP‐1R‐INDEPENDENT PATHWAY IN MICE

The work presented in this chapter has been modified from the following publication: “Activation of Enteroendocrine Membrane Progesterone Receptors Promotes Incretin Secretion and Improves Glucose Tolerance in Mice” Grace B. Flock, Xiemin Cao, Marlena Maziarz, and Daniel J. Drucker. Diabetes August 2012, in press

Author contributions: Xiemin Cao contributed to experiments designed to confirm microarray results (Figure 3.1A) and to the measurement of glucagon levels (Figure 3.10C). Marlena Maziarz contributed with bioinformatics based microarray analysis.

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4.1 ABSTRACT GLP‐1 secretion is principally regulated by nutrients; however, mechanisms regulating the control of GLP‐1 secretion remain incompletely understood. We now demonstrate that GLUTag enteroendocrine cells but not islet ‐cells express a functional progesterone receptor. Progesterone (P4) increased GLP‐1 secretion and ERK1/2 phosphorylation in GLUTag cells, via mechanisms sensitive to the mitogen activated protein kinase inhibitor U0126. The stimulatory effects of progesterone or the synthetic progestin R5020 on ERK1/2 phosphorylation were independent of the progesterone receptor antagonist RU486, whereas a cell impermeable bovine serum albumin‐progesterone conjugate (BSA‐P4) rapidly increased both ERK1/2 phosphorylation and GLP‐1 secretion. Knockdown of progesterone receptor RNA transcripts did not attenuate the stimulatory effects of progestins on GLP‐1 secretion. In contrast, knockdown of the membrane progesterone receptors Paqr5 or Paqr7 eliminated the ability of progesterone to enhance GLP‐1 secretion in GLUTag cells. Enteral P4 administration improved oral glucose tolerance, and increased plasma GLP‐1 and insulin levels in a RU486‐insensitve manner in mice. Furthermore, P4 increased GIP levels following GTT. Unexpectedly however, enteral progesterone improved glucose excursion and increased plasma insulin levels in Glp1r‐/‐ and double incretin receptor knockout (DIRKO) mice. Furthermore, intraperitoneal administration of P4 failed to increase GLP‐1 levels and control glucose excursions during OGTT despite an increase in GIP and insulin levels. These findings extend our concepts of the control of GLP‐1 secretion to include membrane progesterone receptors; however, progesterone does not require a functional GLP‐1R for control of oral glucose tolerance.

4.2 INTRODUCTION Glucagon‐like peptide‐1 (GLP‐1) is a 30 amino acid proglucagon derived peptide (PGDP) secreted from gut endocrine cells that regulates glucose homeostasis by augmenting ‐cell and inhibiting ‐cell function [461]. Conversely, glucagon, a proglucagon‐derived peptide (PGDP) secreted from islet ‐cells plays a key role in the control of blood glucose via regulation of hepatic glucose production [521]. Glucagon excess in the setting of insulin deficiency or resistance contributes to the pathophysiology of diabetes mellitus, whereas defective glucagon secretion leads to an increased risk of hypoglycemia in diabetic subjects. Hence, understanding

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the control of PGDP biosynthesis and secretion in health and disease has direct relevance for the treatment of diabetes mellitus. Enteroendocrine L cells are dispersed as rare single cells scattered throughout the intestinal mucosal epithelium. Although studies of intestinal proglucagon gene expression and PGDP secretion have been carried out using primary intestinal cell cultures [179‐182, 193], L cells often represent less than 1% of the cells in the culture system. Hence, experiments examining intestinal proglucagon gene expression and PGDP synthesis and secretion commonly use immortalized endocrine cell lines [187, 195, 522]. Similarly, experimental approaches for studying islet glucagon biology are limited by the difficulty in obtaining sufficient numbers of pure ‐cells for studies of gene transcription and glucagon secretion. The majority of islet cells in culture systems are ‐cells, with ‐cells estimated to represent less than 10% of islet endocrine cells. Accordingly, many experiments examining glucagon secretion and proglucagon gene expression in ‐cells have utilized immortalized rodent islet cell lines [523, 524]. Enteroendocrine L cells and islet ‐cells both exhibit nutrient‐sensitive regulation of proglucagon gene expression and PGDP secretion; however, the mechanisms controlling proglucagon gene transcription and PGDP secretion in these two cell types have diverged. Nutrients and insulin inhibit islet proglucagon gene expression and glucagon secretion [525]; in contrast, nutrients stimulate proglucagon gene expression and enhance PGDP secretion from gut endocrine cells [207, 526, 527]. To identify mechanisms regulating proglucagon biosynthesis in enteroendocrine cells, we carried out gene expression profiling of enteroendocrine GLUTag cells and for comparative purposes we used islet TC1 cells to search for differentially expressed gene candidates that potentially control PGDP biosynthesis and secretion in gut enteroendocrine L‐cells [190, 528]. Using this approach, we previously identified and characterized a series of ion channels differentially expressed in GLUTag vs TC1 cells [190]. We now have demonstrated that intestinal GLUTag cells but not islet αTC1 cells, express a functional progesterone receptor (PR) and the membrane‐bound progesterone receptors Paqr5 and Paqr7. Progesterone increased levels of proglucagon mRNA transcripts and directly increased GLP‐1 secretion from GLUTag cells in vitro. The progesterone‐dependent stimulation of GLP‐1 secretion was rapid, associated with ERK1/2 activation, insensitive to the classical PR antagonist RU486, and appears mediated by membrane‐bound progesterone receptors.

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Notably, enteral progesterone administration increased plasma GLP‐1, GIP and insulin levels and improved oral glucose tolerance in mice. Unexpectedly however, the actions of progesterone to enhance glucose tolerance were independent of the GLP‐1R and maintained in Glp1r‐/‐ mice. Interestingly, ip administration of progesterone failed to improve glucose homeostasis or increase GLP‐1 levels during OGTT in mice. These findings extend the control of GLP‐1 secretion to encompass sex steroids, and may have implications for understanding how the entero‐insular axis adapts to hormonal changes associated with the metabolic stress of pregnancy.

4.3 RESEARCH DESIGN AND METHODS Tissue culture medium was from Hyclone (Logan, UT), and fetal calf serum (FCS) was from Invitrogen Life Technologies (Burlington, Canada). Antibiotics and chemicals were from Sigma Chemical Co. (St. Louis, MO). Forskolin, progesterone (P4) and RU486 were from Sigma Chemical (St. Louis, MO). R5020, and bovine serum albumin (BSA)‐progesterone (BSA‐P4) were from Steraloids Inc., Newport, RI. Kinase inhibitors were from Calbiochem. Insulin (Humulin R) was from Eli Lilly, Toronto, Ontario, Canada). Antibodies, unless specified otherwise, were from Cell Signaling Technology Ont. CA. Proteinase inhibitor cocktail, phosphatase inhibitors, BSA free of fatty acids (BSA‐FA free) and Tri reagent were from Sigma Chemical (St. Louis, MO). siRNAs were from Invitrogen‐Applied Biosystems (silencer select pre‐designed siRNA). Ultrasensitive mouse insulin ELISA was from Alpco Diagnostics, Salem, NH; mouse/rat total GLP‐ 1 assay kit was from Mesoscale Discovery, Gaithersburg, MD; rat/mouse total GIP ELISA kit was from Millipore, Billerica, MA and glucagon ELISA kit was from Millipore.

4.4.1 Cell culture and microarray experiments For microarray experiments islet αTC1 [529], enteroendocrine GLUTag [530] and SV40‐ transformed mouse fibroblasts, SVT2 (obtained from the American Type Culture Collection) cell lines were maintained in DMEM (4.5 g glucose per litre) supplemented with 15 % fetal calf serum (FCS). Cells were grown to ~80 % confluence and 12 hrs before RNA preparation, media was replaced with fresh media. RNA was prepared with Tri Reagent (Sigma), and purified using the RNeasy Kit (Qiagen). Each single RNA sample submitted for microarray analysis represented

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a pool of three independent plates of cells grown under identical conditions. Microarray analysis was carried out using an n=3 separate pooled RNA samples for each cell line using the Affymetrix MG_U74Av2 Gene Chip. Microarray analysis and expression quantification was performed by GCRMA from Bioconductor (www.bioconductor.org) as described previously [190, 528]. For all other experiments, islet αTC1 [529] and enteroendocrine GLUTag [530] cell lines were maintained, unless otherwise specified, in DMEM (4.5 g glucose per litre) supplemented with 10 % fetal calf serum (FCS).

4.4.2 RNA isolation and analysis For reverse transcription‐polymerase chain reaction (RT‐PCR), first strand cDNA synthesis was generated from total RNA isolated from jejunum and colon of age‐matched males and females C57Bl/6 mice and from islet TC1 and the enteroendocrine GLUTag cell lines using the SuperScript Preamplification System from Invitrogen. Target cDNA was amplified using synthesized specific oligonucleotide forward‐reverse primer pairs and PCR products were probed with an internal 32P‐labeled oligonucleotide. Sequences are provided in table 4.1. Quantitative PCR (qPCR) was performed using Taq‐man assays on demand and ABI PRISM 7900HT (PE Applied Biosystems, Foster City, CA) for proglucagon (Mm00801712); tubulin (Mm00846967); 18S (Hs99999901); progesterone receptor (PR, Mm01176082); progestin and adipoQ receptor family member V (Paqr5, Mm01170057); and progestin and adipoQ receptor family member VII (Paqr7, Mm00910958). Note: SVT2 cell line was used as a model of non‐ endocrine cell for comparative purposes.

Table 4.1 Sequences for primers used to confirm microarray results Gene Symbol Forwerd 5'-3' Reverse 5'-3' Probe Glutamate receptor, ionotropic, kainate 1 Grik1 CTCTCATGCGGCAAGGATC GGCTTTCTGTTTTGCTCCC CGTCCTTTCTGTGTTTGTAGC Bombesin-like receptor 3 Brs3 GAATCCCGGAAGAGAATTGC CCTTCTTGGCACTACTGCC GTAATTCCTGCGTGAACCCC Potassium voltage-gated channel, Kcnd2 GACTCTGTGGCCCTTTGAC CCCCATGAGAAACACTGTG GTGTGGACTGAAGGAAACCA Progesterone receptor PR GGTCGTACAAGCATGTCAG GGATCTTGGGCAACTGGG CTTACCATGTGGCAAATCCC Protein tyrosine phosphatase, non- receptor type 5 Ptpn5 CCCCATCTATTGTCCTGGC GGTCTCTGCCATCCACATC GCCAGGAAAGGAGCACTGAA Cathepsin C Ctsc CCTTTCAACCCCTTCGAGC GACAACTGAACCACTGCTC CGGGTCTTTATCACTCACAG Caudal type homeo box 2 Cdx2 CAGAACCGCAGAGCCAAG CTTCGTTTGTCGTTGCTGG CTGAGCCATGAGGAGTATGG Unc4.1 homeobox (C. elegans) Uncx4.1 CTTTCCGGGACCTAACAGC CCATCGACACAGCGTTTTC GATGCGCTTAGCCAGTCGAG T-cell acute lymphocytic leukemia 1 Tal1 CCAAGGGCACAGCAACTAG GTACAAAGTCCAGGCCCC CATAGCACGCCATGTCTGTG

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4.4.3 Cell transfection GLUTag cells were grown to 80% confluence in DMEM high glucose (4.5 g glucose per litre) supplemented with 10% fetal calf serum FCS and transfected with [‐300]GLU‐Luc, [531] or MMTV‐Luc (positive control) using Fugene (Roche Applied Science, Quebec, CA) or lipofectamine (Invitrogen) as indicated, following the manufacturer’s protocol. 24 hrs after transfection the media was replaced with DMEM high glucose supplemented with 0.5% FCS and 12 hr later cells were treated over night with R5020 (20 nM), progesterone (P4) (20nM), forskolin (20 M), dexamethasone (10‐7 M), DMSO, or ethanol. Cells were then harvested for analysis of luciferase activity as described previously [18, 531]. Luciferase values were normalized to total protein content determined by a BCA protein assay kit (Pierce, IL; USA). For knockdown of RNA transcripts GLUTag cells were grown to 50 % cell density and siRNA was transfected using lipofectamine RNAiMAX (Invitrogen) following the manufacturer's protocol. siRNAs included the nuclear progesterone receptor (si‐PR) (s71548 plus s71549), the progestin and adipoQ receptor family member V (si‐Paqr5) (s92240), the progestin and adipoQ receptor family member VII (si‐Paqr7) (s163230 ). The negative control (si‐Neg. Ctrl.) was from Albion, cat# 4390844. Efficiency of RNA knockdown was assessed by quantitative polymerase chain reaction. Concomitant quantification of GAPDH mRNA transcripts was used to normalize mRNA transcript levels.

4.4.4 Signal transduction studies GLUTag cells were incubated in DMEM high glucose (4.5 g glucose per litre) supplemented with 10% FCS. Twelve hours before treatment, cells were washed twice with PBS and media was replaced with DMEM high glucose supplemented with 0.5 % FCS. Cells were then washed twice with PBS and incubated in DMEM without FCS for 2 hrs followed by treatment for 10 or 30 min with the progestin R5020 (20nM), progesterone (P4) (20nM), insulin (10nM), epidermal growth factor (EGF) (25 ng/ml), covalently bound (BSA‐P4; 20nM progesterone) or ethanol (vehicle control). Whole cell extracts were prepared in radioimmunoprecipitation assay buffer (RIPA) (Tris 50 mM, NaCl 150 mM, SDS 0.1 % , Na‐Deoxycholate 0.5 % ,Triton X 100 1% ) containing proteinase inhibitor cocktail and phosphatase inhibitor. Protein content was measured by BCA protein assay kit (Pierce, IL; USA) and 40 ug of total protein was separated by

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electrophoresis in a 10 % SDS PAGE gel. Following transfer to a nylon membrane, blots were probed with antisera directed against Phospho‐P44/42 MAP Kinase (pERK1/2), P‐Creb (Ser133), P‐AKT (Ser473) and AKT. HSP90 was used as loading control. In separate experiments, cells were pre‐treated for 30 min with H89 (10uM), UO126 (10uM), or PD98059 (50uM), or for 15 min with the classical nuclear progesterone receptor antagonist RU486 (1uM) followed by R5020 (20nM), P4 (20nM) or vehicle. Densitometry analysis was done with a Kodak image station 4000 MM PRO and values were normalized to levels of HSP90 in the same experiment. cAMP was measured from dried aliquots of ethanol cell extracts using a cAMP radioimmunoassay kit (Biomedical Technologies, Stoughton, MA).

4.4.5 GLP‐1 levels GLUTag cells grown to 80 % confluence in DMEM with 10% FCS were washed with PBS and pre‐incubated for 2 hours in DMEM without FCS. Media was replaced with fresh DMEM high glucose (4.5 g glucose per liter) without FCS supplemented with either R5020 (20 nM), P4 (20nM), BSA‐P4 (20nM of P4), forskolin (10 µM) or 10 % FCS for 2h. Media and cell extracts were collected in the presence of proteinase inhibitor cocktail and GLP‐1 levels were measured using the total GLP‐1 RIA (Millipore Corporation, MA) or the total GLP‐1 assay kit (Mesoscale Discovery, Gaithersburg, MD). Total GLP‐1 secreted into the media was normalized to total protein content or total GLP‐1 (cell plus media) content.

4.4.6 Animal experiments Mouse experiments were carried out according to protocols approved by the Mt. Sinai Hospital and the Toronto Centre for Phenogenomics (TCP) Animal Care Committees.

Glucose Tolerance Tests Age‐matched male mice were fasted for 4 hours, and a single oral dose of vehicle (V) (80% PEG400, 10% Tween 80, 10% ethanol) (day 1) or progesterone (P4) (100 µg per mouse) (day 2) was administered 15 min prior to oral or ip glucose administration (1.5 g/kg body weight). In a separate experiment, mice were given a single oral dose of either V (80% PEG400, 10% Tween 80, and 10% ethanol) or the classical nuclear progesterone receptor antagonist RU486 (100 mg/kg) as described [532]. Thirty minutes later mice received a single oral dose of V (80%

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PEG400, 10% Tween 80, 10% ethanol) or P4 (100 µg / mouse) followed by an oral glucose load (1.5 g/kg body weight) 15 min later. Blood glucose levels were measured by sampling from the tail vein of gently held conscious mice, from 5‐90 min after glucose administration. At the 5‐min time point, a blood sample (150 µl) was collected and immediately mixed with 15 µl of a chilled solution containing 5000 kIU/ml Trasylol (Bayer, Toronto, ON), 32 mM EDTA, and 0.01 mM Diprotin A (Sigma St. Louis, MO) for measurement of insulin, GLP‐1 and GIP levels. When P4 was administered intraperitoneally (ip), mice were fasted for four hours and were given a single ip dose of either corn oil (vehicle) (day 1) or P4 (100ug) (day 2) 15 min prior to oral glucose load. A blood sample was collected 5 min after glucose for the assessment of total GLP‐1, total GIP and glucagon. A second blood sample was obtained at 15 min after glucose administration for the measurement of insulin and glucagon. Plasma was obtained by centrifugation at 4 C and stored at ‐80 C.

Gastric emptying Liquid‐phase gastric emptying was assessed using the acetaminophen absorption test [469]. C57BL/6 and DIRKO male mice, 10‐12 weeks of age were fasted for 4 hours and given a single dose of either V (80% PEG400, 10% Tween 80, 10% ethanol) (day 1) or P4 (100µg/mouse) (day 2) 15 min before oral administration of a solution of glucose 15% and acetaminophen 1% (Sigma, St Louis,MO ) at a dose of 1.5 g/kg glucose‐0.1 g/kg acetaminophen. Tail vein blood (50 µl) was collected into heparinized tubes at 15 and 30 min after glucose/acetaminophen administration. Plasma was separated by centrifugation at 4 C and stored at –20 C until measurement of acetaminophen levels using an enzymatic‐spectrophotometric assay (Diagnostic Chemicals Ltd., Oxford, CT).

Insulin Tolerance Test Ten‐week‐old age‐matched C57BL/6 and DIRKO male mice were fasted for 5 hrs. A single oral dose of V (80% PEG400, 10% Tween 80, 10% ethanol) (day 1) and P4 (100 µg/mouse) (day 2) was given 15 min prior to the administration of 1.2 U / kg of insulin (Humulin R, Eli Lilly, On. Canada). Blood glucose was determined at 0, 15, 30, 60, 120 and 180 minutes.

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Proglucagon expression study after P4 administration Ten‐week‐old C57BL/6 male mice were given regular chow supplemented with P4 (RC‐P4) or vehicle (corn oil) (RC‐V) for 48 hrs. Diet was prepared as follow: 100 grams of regular chow were grinded and blended with 10 ml corn oil (vehicle) or 0.33 mg progesterone previously dissolved in 10 ml of corn oil. The powdered diet was then kept moist by addition of water to achieve paste consistency. Daily estimated dose of P4 was 10 mg/day/mouse. Mice were then euthanized and jejunum and colon were flashed cleaned with cold PBS and collected for RNA preparation. Real time PCR was performed as described above using the assay on demand for proglucagon (Mm00801712, Applied Biosystems). Blood was collected by cardiac puncture and immediately mixed with a chilled solution containing 5000 kIU/ml Trasylol (Bayer, Toronto, ON), 32 mM EDTA, and 0.01 mM Diprotin A (Sigma St. Louis, MO). Plasma was obtained by centrifugation at 4 C and stored at –80 C until determination of total GLP‐1 using mouse/rat total GLP‐1 assay kit (Mesoscale Discovery, Gaithersburg, MD) and progesterone using enzyme‐ linked immunosorbent assay kit (Neogen, Lexington, Ky).

4.4.7 Statistical analysis Statistical significance was assessed by one‐way ANOVA using Bonferroni multiple comparison post hoc test and, where appropriate by paired Student’s t test using GraphPad Prism 4 (Graph‐ Pad Software, San Diego, CA). A P value of <0.05 was considered to be statistically significant.

4.4 RESULTS Differential expression of genes in glucagon‐producing cell lines We identified several mRNA transcripts differentially expressed in GLUTag cells compared to TC1 cells (Figure 4.1A). For example, the transcription factors Cdx2, Unc4.1, and Tal1, were preferentially expressed in GLUTag cells (Figure 4.1). Similarly, mRNA transcripts for the ion channel Kcnd2, the glutamate receptor Grik1, and the protein tyrosine phosphatase Ptpn5 were detected in RNA from GLUTag, but not TC1 cells. Unexpectedly, we also detected robust expression of the classical progesterone receptor (PR), in GLUTag, but not TC1 cells (Figure 4.1A).

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PR‐mediated regulation of proglucagon gene expression in GLUTag cells We next ascertained whether the GLUTag PR mRNA transcript gives rise to a transcriptionally competent PR. GLUTag cells were transfected with the PR‐responsive MMTV‐ Luc reporter plasmid and treated with the synthetic progestin R5020, or the glucocorticoid dexamethasone. R5020 and to a lesser extent dexamethasone significantly activated MMTV‐Luc activity in GLUTag cells (Figure 4.2A). In contrast, R5020 had no effect on MMTV‐Luc activity in TC1 cells; however, dexamethasone robustly induced MMTV‐Luc activity in the same experiments (Figure 4.2B). R5020 also significantly increased the levels of proglucagon mRNA transcripts in GLUTag cells when incubated in the presence of 0.1% FCS (Figure 4.2C) but failed to do so in the presence of charcoal stripped FCS (data not shown), thus other factors present in the serum are required for R5020‐regulation of proglucagon gene transcription. Furthermore, standard chow diet (RC) supplemented with P4, failed to elevate proglucagon mRNA transcript levels in jejunum and colon of male mice (Figure 4.2D). Surprisingly, an almost significant (p=0.0508) increase in circulating GLP‐1 levels associated with a significant increase in P4 levels was seen in RC‐P4 compared to RC‐V treated mice (Figure 4.2E). Furthermore, although forskolin significantly increased the transcriptional activity of ‐ [300]GLU‐Luc, a cAMP‐ regulated proglucagon promoter plasmid harboring the G1‐G5 promoter elements [533], R5020 had no effect on proglucagon promoter‐dependent luciferase activity in transfected GLUTag cells (Figure 4.2F). However, R5020, but not forskolin significantly increased MMTV‐Luc activity in the same experiments (Figure 4.2F).

Progesterone mediated GLP‐1 secretion and signaling pathways As progesterone regulates hormone secretion through both genomic and non‐genomic mechanisms [420, 534], we examined levels of secreted GLP‐1 in GLUTag cells. R5020 significantly stimulated GLP‐1 secretion from GLUTag cells (Figure 4.3A). In contrast to forskolin, which produced a robust and sustained activation of cyclic adenosine monophosphate response element‐binding (CREB) phosphorylation (Figure 4.3B), R5020 produced a biphasic effect, with increased levels of phospho‐CREB detected at 5 minutes followed by a secondary increase at 30‐60 minutes (Figure 4.3B). R5020 rapidly increased ERK1/2 (Figure 4.3C) but not AKT phosphorylation (Figure 4.3D).

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TC 1 GLUTag SVT2 Gene Glutamate receptor, ionotropic, kainate 1 (Grik1) Bombesin‐like receptor 3 (Brs3) Potassium voltage‐gated channel, Shal‐related family, member 2 (Kcnd2) Progesterone receptor (PR) Protein tyrosine phosphatase, non‐receptor type 5 (Ptpn5) Cathepsin C (Ctsc) Caudal type homeobox 2 (Cdx2) Unc4.1 homeobox (C. elegans) (Uncx4.1) T‐cell acute lymphocytic leukemia 1 (Tal1)

Figure 4.1 Progesterone receptor is differentially expressed in GLUTag vs αTC1 cells. Examples of genes differentially expressed in enteroendocrine GLUTag cells compared to TC1 cells. SVT2 cells were used as a model of non‐endocrine cell line (negative control). Expression of selected genes identified by microarray analysis was assessed by RT‐PCR as described in Methods. SVT2 cells were used as a control SV40 T antigen transformed non‐endocrine cell line. The expression of the indicated genes was assessed by RT‐PCR. PCR products were probed with an internal 32P‐labeled oligonucleotide.

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5 A GLUTag D MMTV-Luc 4 3 *** MMTV-Luc + R5020 *** MMTV-Luc + Dex 3 2 2 RLU 1 1 relative expression Proglucagon mRNA MMTV-Luc

0 0 J-V J-P4 C-V C-P4

B E 8 *** 40 ** 30 p = 0.0508 6 TC1 30 20 4 20 RLU 10 P4 (ng/ml) MMTV-Luc 2 10 tGLP1 (pg/ml)

0 0 0 V P4 V P4

C F

4 4 4 ** 0.1% FCS ** *** R5020 3 3 3 10% FCS Bas al R5020 2 2 2 RLU RLU Forskolin 1 1 1 Relative expression Relative Proglucagon mRNA 0 0 0 [- 300]GLU-Luc MMTV-Luc

Figure 4.2 The PR expressed in GLUTag cells is functional and transactivates the MMTV promoter; however, P4 does not modulate proglucagon gene transcription in vivo. (A) GLUTag and (B) αTC1 cells transfected with MMTV‐Luc were treated with R5020 (20 nM) or vehicle (V‐ethanol) as described in Methods. Luciferase activity was assessed 12 hours after treatment. Dexamethasone (Dex) (10‐7M) was used as a positive control. Results depict mean ± SD from two independent experiments each one done in quadruplicate and are expressed relative to the luciferase activity measured for vehicle‐treated cells. *** p<0.001 vs V‐treated cells. (C) The progestin R5020 (20 nM) increases levels of proglucagon mRNA transcripts in GLUTag cells. Proglucagon mRNA levels were assessed by qPCR following incubation of the cells with R5020 (20 nM) or V for 48 hrs in the presence of 0.1% FCS. Data are mean ± SD of two independent experiments and are expressed relative to V‐treated cells. ** p<0.01 vs. V‐treated cells. (D) Enteral P4 does not modulate intestinal proglucagon mRNA levels in vivo. Male mice (n=4 per group) were fed a regular chow diet supplemented with P4 (daily estimated dose of P4: 10 mg/day) or V (corn oil) for 48 hrs. Jejunum (J) and colon (C) were collected for mRNA analysis by qPCR. Results are expressed relative to the values for J‐V. (E) Blood was collected from mice used in (D) for the assessment of postprandial circulating P4 and total GLP‐1 (tGLP‐1). ** p<0.01, *** p<0.001 vs. V‐treated mice. (F) Transcriptional activity of proglucagon promoter and MMTV‐Luc plasmids in GLUTag cells. GLUTag cells transfected with [‐300] GLU‐Luc, and MMTV‐Luc (positive control) and treated with R5020 (20nM), Forskolin (20 M) and vehicle (V) as described in Methods. Results are expressed relative to basal conditions (V‐treated cells). Vehicle treatment resulted in no significant differences in luciferase activity relative to basal conditions (not shown). * p<0.05, ** p<0.01, *** p<0.001.

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A E 40 15 *** 30 *** 10 20 tGLP-1 tGLP-1 5 * 10 * (relative level) (relative (relative level) (relative 0 0 0 6 2 2 lin al 20 in o s 0 ol k Basal R50 Ba R5 sk UO126 UO1 + ors 0 F For 2 0 R5 B F

Basal Forskolin R5020 4 * R5020 3 0’ 5’ 10’ 20’ 30’ 60’ 5’ 10’ 20’ 30’ 60’ Forsk pCREB 2 HSP90 1

0 C Relative cAMP levels 30 60 R5020 Vehicle Time (min)

0’ 10’ 30’ 10’ 30’ pERK1/2

HSP90

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Figure 4.3 Progesterone stimulates GLP‐1 secretion in a MAPK‐dependent manner in GLUTag cells. (A) GLP‐1 secretion in GLUTag cells treated with R5020. Total GLP‐1 secreted in the media (pg/ml) over 2 hrs was normalized to total cell protein content (µg). Results depict mean ± SD from two independent experiments performed each in triplicate and are expressed relative to the levels of GLP‐1 secreted under basal conditions (V‐ treated cells). (B ‐ D) Western blot analysis of (B) CREB phosphorylation and (C) ERK1/2 phosphorylation and (D) AKT phosphorylation in GLUTag cells treated with R5020 (20 nM). Anti‐heat‐shock protein 90 (HSP 90) was used to monitor loading and transfer conditions. (E) R5020 (20 nM) stimulates GLP‐1 secretion in a U0126‐sensitive manner in GLUTag cells. GLUTag cells were pre‐treated for 30 min with U0126 (10 M) or V (DMSO) followed by treatment with R5020 (20nM) or V (ethanol). Total GLP‐1 secreted in the media (pg/ml) over 2hrs was normalized by protein cell content (µg) and expressed relative to basal conditions (V‐treated cells). (F) R5020 (20 nM) does not increase levels of cAMP. GLUTag cells were incubated as described in Methods and treated for 30 min or 60 min with R5020 (20nM) or Forskolin (100 M). cAMP was measured from dried aliquots of ethanol cell extracts. Results shown are the mean ± SD of a representative experiment done in quadruplicate. * p<0.05, ** p<0.01, *** p<0.001 vs. V‐treated cells.

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Consistent with the importance of ERK1/2 as downstream effectors of progestin action in GLUTag cells, co‐treatment with UO126, an inhibitor of both active and inactive MEK1,2 eliminated the R5020‐mediated induction of GLP‐1 secretion (Figure 4.3E). Similarly PD98059, an inhibitor of active MEK1,2, prevented P4‐induced ERK1/2 phosphorylation (data not shown). Unlike forskolin, which also stimulates GLP‐1 secretion, R5020 had no effect on cAMP accumulation in GLUTag cells (Figure 4.3F).

Progesterone mediated GLP‐1 secretion: genomic vs. non‐genomic As the secretory activity of the enteroendocrine L cell is sensitive to ambient glucose levels [186], we examined whether progesterone actions in GLUTag cells were glucose‐dependent. Both progesterone and R5020 modestly augmented ERK1/2 phosphorylation in the absence of glucose; however, the induction of ERK1/2 phosphorylation was considerably greater in cells incubated at 25 mM glucose (Figure 4.4A). Unexpectedly, R5020 and progesterone both increased ERK1/2 phosphorylation in the presence of the classical PR antagonist RU486 (Figure 4.4B). Accordingly, we hypothesized that the actions of progesterone in GLUTag cells may be mediated through non‐genomic mechanisms. We detected expression of mRNA transcripts for not only the progesterone receptor, but also the progestin and adipoQ receptor family member V (Paqr5), and the progestin and adipoQ receptor family member VII (Paqr7) in RNA isolated from the murine jejunum and colon, as well as GLUTag cells (Figure 4.4C). For comparative purposes, levels of

proglucagon mRNA transcripts were assessed using the same aliquots of RNA (Figure 4.4C).

Membrane‐bound progesterone receptor and GLP‐1 secretion modulation To further examine the hypothesis that membrane‐localized actions of progesterone activate ERK1/2 phosphorylation and GLP‐1 secretion in GLUTag cells, we utilized a progesterone‐albumin covalently linked molecule that is unable to penetrate the cell membrane (BSA‐P4) [535]. BSA‐P4 (20 nM of P4) rapidly and significantly increased ERK1/2 phosphorylation to levels comparable to that achieved with R5020 (20nM) or progesterone (20nM) (Figure 4.4D). Similarly, BSA‐P4 (20 nM of P4) significantly increased GLP‐1 secretion from GLUTag cells to levels comparable to that seen with R5020 or P4 (20 nM) (Figure 4.4E).

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A B

No glucose 25 mM glucose P4R5020 P4 R5020 0’ 10’ 30’ 10’ 30’ 0’ 10’30’ 10’ 30’ pERK1/2 HSP90

C

6 5000 50 3020 5 3000 30 *** 1520 4 1000 10 20 2 4 3 4

PR *** Paqr7 3

Paqr5 2 * * 1 2 1 2 Proglucagon 1 0 0 0 0 (relative expression) (relative (relative expression) (relative (relative expression) (relative ) ) 1 ) ) ) g 1 ) 1 C (F (F ag C1 (M) C (F) ag C (F) (F) T (M) T T J (F) T (M) T J J (M C C (M  J J (M C C U a J C (F) C (M) UTa  J J (M) C (F C U aT L L GLUTag GL G G

D E F ### ** *** 3 2 *** 15 * *** * *** V * 10 2 R5020 5 BSA-P4 1 3 BSA

1 (RLU) tGLP-1 2 MMTV-Luc 1 phosphorilation (relative levels) 0 0 0

Relative level of pERK1/2 P4 P4 Basal R5020 BSA-P4 Basal R5020 BSA-P4

Figure 4.4 Progesterone stimulates GLP‐1 secretion via non‐genomic mechanisms activating Paqr5 and Paqr7 in GLUTag cells (A) Western blot analysis of ERK1/2 activation in cells incubated in DMEM without glucose vs. DMEM high glucose and treated with P4 (20 nM) or R5020 (20 nM). (B) P4‐stimulated ERK1/2 phosphorylation is not inhibited by RU486 in GLUTag cells. Western blot analysis of ERK1/2 phosphorylation in cells pre‐treated for 15 min with the PR antagonist RU486 (1M) followed by treatment for 10 or 30 min with R5020 (20nM), P4 (20nM) and epidermal growth factor (EGF) (25 ng/ml, positive control for ERK1/2 phosphorylation). Anti‐heat‐shock protein 90 (HSP 90) was used to monitor loading and transfer conditions. (C) Expression of PR, Paqr5, Paqr7 and proglucagon (Gcg) mRNA transcripts in the jejunum (J) and colon (C) of female (F) and male (M) C57Bl/6 mice (n=5 per group) as assessed by qPCR. Levels of mRNA transcripts are expressed relative to the expression in the jejunum of female mice. (D) Covalently bound BSA‐progesterone conjugate (BSA‐P4) increases ERK1/2 phosphorylation. GLUTag cells were treated for 10 min with the progestin R5020 (20nM), progesterone (P4) (20nM) or covalently bound (BSA‐P4; 20nM progesterone). Cells treated with vehicle (ethanol) plus BSA free of fatty acids (20nM) were used as the basal control. Figure shows the densitometry analysis of ERK1/2 phosphorylation normalized to HSP90 in the same experiment and expressed relative to basal conditions. Results depict mean ± SD of two independent experiments each done in triplicate. * p<0.05, ** p<0.01, *** p<0.001. (E) BSA‐P4 (20 nM progesterone) stimulates GLP‐1 secretion in GLUTag cells. Cells were treated for 2 hrs with either R5020 (20 nM), P4 (20nM) or BSA‐P4 (20nM of P4). Cells treated with vehicle (ethanol) plus BSA free of fatty acids (20nM) was used as the basal control. Total GLP‐1 secreted in the media (2 hr incubation) (pg/ml) was normalized to total cell‐protein content (g). Results depict mean ± SD of two independent experiments, each performed in quadruplicate and expressed relative to levels of GLP‐1 secreted under basal conditions. * p<0.05, ** p<0.01*** p<0.001 (F) BSA‐P4 (20 nM progesterone) does not transactivate the MMTV‐Luc promoter. GLUTag cells transfected with the MMTV‐Luc were treated for 12 hrs with R5020 (20 nM), BSA‐P4 (20 nM of P4), V or BSA‐free of fatty acids (BSA) (20 nM). Results are expressed relative to the luciferase activity in V treated cells and represent the mean and SD of a representative experiment done in triplicate. *** p<0.001.

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In contrast, BSA‐P4, unlike R5020, had no effect on the activity of MMTV‐Luc in transfected

GLUTag cells (Figure 4.4F). Interestingly, BSA‐P4 increased GLP‐1 secretion to a greater extent than P4 in GLUTag cells (Figure 4.4E). To identify the receptor(s) required for progesterone‐mediated regulation of GLP‐1 secretion we used siRNA to reduce expression of the classical PR, the membrane‐bound progestin and adipoQ receptor family member V (Paqr5) or the membrane‐bound progestin and adipoQ receptor family member 7 (Paqr7) (Figure 4.5A). A reduction of PR mRNA by more than 50% did not impair and actually enhanced the stimulatory effects of P4 on GLP‐1 secretion (Figure 4.5B). However, reduction in levels of Paqr5 and Paqr7, alone or together, abrogated the ability of P4 to stimulate GLP‐1 secretion (Figure 4.5B). The same results were observed using R5020 (not shown). Hence, progesterone stimulates GLP‐ 1 secretion via activation of both Paqr5 and Paqr7 independent of the PR in GLUTag cells.

Enteral progesterone regulates glucose‐stimulated GLP‐1 secretion in vivo We then examined whether progesterone increases plasma levels of GLP‐1 and lowers glucose in mice. A single dose of oral progesterone significantly improved oral glucose tolerance, in association with increased plasma levels of insulin and total GLP‐1 immunoreactivity in mice (Figure 4.6A‐D). Consistent with data obtained in GLUTag cells, the progesterone‐mediated improvement in glucose tolerance (Figure 4.7A, B) and increase in plasma levels of GLP‐1 (Figure 4.7C) was RU486‐independent.

Enteral progesterone controls glucose homeostasis independent of incretin signaling To assess whether progesterone‐mediated control of glucose tolerance requires the presence of a functional GLP‐1R we treated Glp1r‐/‐ and littermate control mice with a single oral dose of progesterone prior to administration of oral glucose. Progesterone improved oral glucose tolerance and significantly increased insulin levels after an oral glucose load in WT and Glp1r‐/‐ mice (Figure 4.8A, B). Hence, although progesterone stimulates GLP‐1 secretion in vitro and increases plasma levels of GLP‐1 in vivo, the actions of progesterone to control glucose homeostasis and increase insulin levels are independent of the GLP‐1R.

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A 1.5 1.5 1.5

1.0 1.0 1.0

PR * *** *** Paqr7 Paqr5 ** 0.5 0.5 0.5 ** (relative expression) (relative (relative expression) (relative (relative expression) (relative 0.0 0.0 0.0 l l l R a trl a P s C qr7 si aqr5 aqr7 aqr7 Basa Ctrl Ba P P Bas iPa P g. si eg s iN Ne s siNeg Ctrl si qr5+si iPa B siPaqr5+si s *** 6 ### 5 V 4 P4 3 * 2 * 1 tGLP-1 (pg/ml) tGLP-1 level) (relative 0 siPaqr5 + siPR Basal

siPaqr7 siPaqr7 siPaqr5 siNeg.Ctrl Figure 4.5 Progesterone stimulates GLP‐1 secretion in GLUTag cells independent of the classical progesterone receptor (PR). (A) siRNA knock down reduces levels of PR, Paqr5, and Paqr7 mRNA transcripts in GLUTag cells. siRNA was transfected as described in Methods and the efficiency of RNA knockdown was assessed by qPCR. Levels of mRNA transcripts after knockdown are expressed relative to the levels detected under basal conditions. * p<0.05, ** p<0.01 vs. basal. (B) Relative GLP‐1 secretion in cells transfected with the indicated siRNAs and treated for 2 hrs with R5020 (20 nM), P4 (20 nM), or V (ethanol). Total GLP‐1 secreted in the media (pg/ml) was normalized to total cell protein content (g). Results are expressed relative to levels of GLP‐1 in cells treated with V and depict mean ± SD from three independent experiments each done in triplicate. * p<0.05, *** p<0.001 vs. basal. ###p<0.001, siPR + P4 vs. basal + P4.

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Tail bleed Tail OralV P4 or Oral glucose -15’ 0’ 5’ 90’ Time (min) A B

22 500 * 19 V P4 16 250 13 10 Glucose (mM) * min) x (mM AUC 7 0 V P4 0 15 30 45 60 75 Time (min)

C D 3 ** 200 * 2

100 1 Insulin (ng/ml) 0 (pg/ml) tGLP-1 0 V P4 V P4

Figure 4.6 Progesterone improves oral glucose tolerance and increases plasma GLP‐1 levels in mice. (A) Plasma glucose excursion. (B) Glucose area under the curve (AUC) for OGTT shown in (A). (C) Glucose stimulated insulin and (D) Glucose stimulated Total GLP‐1 (tGLP‐1) in plasma obtained at the 5 minute time point of the OGTT shown in (A). V=vehicle (dashed line), P4=progesterone (solid line). C57BL/6 mice were fasted and given vehicle or P4 (100 µg) 15 min before oral glucose administration followed by tail bleed, 5 min after glucose loading for the measurement of insulin and tGLP‐1. (n = 9 mice). * p<0.05, ** p<0.01.

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Oral RU486 or V V or P4 Oral Oral glucose bleed tail

-75’ -15’ 0’ 5’ 90’ Time (min)

A 25 V P4 20 RU486 RU486-P4 15 10 5 Glucose (mM) Glucose 0 0 15 30 45 60 75 90 B Time (min)

600 ** V 500 * P4 400 RU486 300 RU486-P4 200 100 AUC (mMAUC min) x 0

C

75 * ** 50

25 tGLP-1 (pM) tGLP-1 0

Figure 4.7 Progesterone lowers glucose and increases plasma GLP‐1 levels in mice via RU486‐insensitive mechanisms. (A) Glucose excursion during an OGTT in mice pre‐treated with the PR antagonist RU486 (100 mg/kg) in the presence or absence of enteral P4 (100 µg) administration. V=vehicle (empty circle‐dashed line), RU486 (empty square‐dashed line), P4=progesterone (solid square‐solid line), (RU486‐P4 (solid circle‐solid line) (B) Area under the curve for glucose levels of OGTT shown in (A) and (C) Total plasma GLP‐1 (tGLP‐1) levels at the 5 minute time point from the OGTT shown in (A). Mice were fasted and given orally vehicle or RU486 (100 mg/kg) 30 min before oral P4 (100 µg) or vehicle administration followed by OGTT 15 min later. Mice were then bled 5 min after oral glucose administration for the measurement of tGLP‐1. (n = 7 mice per group) * p<0.05; ** p<0.01.

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Oral P4 or V P4 or Oral Oral glucose bleed Tail -15’ 0’ 5’ 90’ Time (min)

AB

1000 1000 25 WT Glp1r-/- 800 ** 800 V 30 600 600 P4 ** 20 * 400 400 *** 200 200 20 AUC (mM x min) 0 15 (mMAUC x min) 0

10 10 Glucose (mM) Glucose Glucose (mM) V 5 P4 0 0 0 10 20 30 40 50 60 70 80 90 100 0 10 20 30 40 50 60 70 80 90 100 Time (min) Time (min)

CD DIRKO 25 Gipr-/- 30 *** 20 *** 20 15 *** 1000 1000 ** 10 800 800 600 V 10 600 Glucose (mM) Glucose Glucose (mM) 5 400 P4 400 200 200 AUC (mMAUC x min) 0 (mMAUC x min) 0 0 0 0 20 40 60 80 100 0 20 40 60 80 100 Time (min) Time (min)

Figure 4.8 Progesterone improves oral glucose tolerance independent of incretin receptors. Glucose excursions and area under the curve (AUC) during OGTT for (A) WT (n = 14 mice), (B) Glp1r‐/‐ (n = 8 mice), (C) Gipr‐/‐ (n=10) and (D) DIRKO mice (n = 17 mice). Age‐matched mice were treated with a single oral dose of progesterone P4 (100 g)) (solid line) or vehicle (V) (dashed line) 15 min before an oral glucose load. * p<0.05, *** p<0.001 vs. vehicle treated mice.

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To interpret this finding we investigated whether enteral progesterone stimulated GIP secretion and regulated glucose homeostasis via Gipr‐dependent mechanisms. We repeated these studies in Gipr‐/‐ and double incretin receptor knockout (DIRKO) mice. P4 failed to improve glucose homeostasis in Gipr‐/‐ mice (Figure 4.8C). Surprisingly, however, P4 significantly reduced glucose excursions in DIRKO mice (Figure 4.8D). Plasma insulin levels measured 5 minutes following glucose administration were significantly increased by P4 in WT and Glp1r‐/‐ (Figure 4.9A). In accordance with the lack of a P4‐mediated enhancement of glucose control in Gipr‐/‐ mice, P4 did not significantly increase insulin levels in these mice (Figure 4.9A). Surprisingly, insulin levels were not increased by P4 in DIRKO mice despite a significant improvement in glucose excursions (Figure 4.9A); however, a subsequent time‐ course analysis revealed that P4 significantly increased circulating levels of insulin at 15 and 20 minutes following oral glucose administration in DIRKO mice (Figure 4.9B). Thus, a time course for Gipr‐/‐ mice is also necessary. Plasma levels of glucose‐stimulated GIP were significantly increased in WT and Gipr‐/‐ mice following P4 administration. A detectable, but non‐significant, increase of glucose‐stimulated plasma GIP was observed in Glp1r‐/‐ mice treated with P4 compared to vehicle‐treated animals. In contrast, a modest, but not significant, increase of GIP levels following an oral glucose load was seen in P4‐treated DIRKO mice (Figure 4.9C). Similarly, plasma levels of GLP‐1 were significantly increased by P4 in Gipr‐/‐ but not in DIRKO mice (Figure 4.9D). Our results demonstrate that P4 regulates GLP‐1 and GIP secretion in vivo; however, abrogation of incretin signaling does not prevent enteral progesterone actions on glucose control. Interestingly, elimination of GIP signaling in the presence of intact GLP‐1 signaling prevents P4‐mediated improvement of glucose excursions. GLP‐1 activation engages multiple mechanisms to improve glucose control. GLP‐1, apart from its insulinotropic actions, inhibits glucagon secretion from islet α‐cells and reduces the rate of gastric emptying, resulting in reduced postprandial glucose excursions [141, 448, 449]. We hypothesized that modulation of the rate of gastric emptying could be another mechanism involved in enteral P4 improvement of glucose homeostasis. To test this possibility, we performed liquid‐phase gastric emptying studies using the acetaminophen absorption test [469] in age‐ matched male C57Bl6 and DIRKO mice.

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Measure blood glucose Tail bleed Oral P4 or V Oral glucose Tail bleed -15’ 0’ 5’ 90’ Time (min) 10’ 15’ 20’ Measure GSI (DIRKO, time course)

Measure: Total GIP, Total GLP-1, Insulin

AB2.5 4 ** 2.0 * * 3 1.5 * 2 p=0.0633 1.0 1

Insulin (ng/ml) 0.5

Insulin (ng/ml) Insulin 0 0.0 WT Glp1r-/- Gipr-/- DIRKO 10 15 20 n=15 n=11 n=10 n=17 Time (min)

C D 1000 V 80 * P4 * 800 60 600 * 40 400 20 200 tGIP (pg/ml) 0 tGLP-1 (pg/ml) 0 WT Glp1r-/- Gipr-/- DIRKO Gipr-/- DIRKO n=14 n=3 n=5 n=7 n=6 n=7

Figure 4.9 Plasma incretin and insulin levels following enteral progesterone administration during OGTT. Age‐matched WT, Glp1r‐/‐, Gipr‐/‐ and DIRKO mice were treated with a single oral dose of progesterone P4 (20 nM) 15 min before an oral glucose load. For A, C and D, a blood sample was collected 5 minutes after oral glucose administration for the assessment of glucose‐stimulated plasma insulin and incretin levels. For B, an independent group of DIRKO mice (n = 10 mice) were treated as described above and bled at the indicated time points following glucose administration for the assessment of plasma insulin levels during an OGTT. * p<0.05, *** p<0.001 vs. vehicle treated mice. (A) GSI, (B) Time course for GSI, (C) Total GIP levels (tGIP) and (D) Total GLP‐1 levels (tGLP‐1). * p<0.05, ** p<0.01, *** p<0.001.

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Our results demonstrate that enteral progesterone does not modify the rate of gastric emptying in mice (Figure 4.10A). To assess whether acute enteral P4 differentially modulates insulin sensitivity in mice lacking incretin signaling compared to WT controls, we administered a single oral dose of V (day 1) or P4 (day 2) to age‐matched C57Bl/6 and DIRKO mice, 15 min prior to an ITT. P4 administration did not result in an enhancement of insulin sensitivity as assessed by ITT (Figure 4.10B, C). Our results demonstrate that acute enteral P4 administration does not modify insulin sensitivity in mice. To elucidate the importance of enteral vs. progesterone exposure for the glucoregulatory actions of P4, we tested whether ip administration of P4 improved glucose excursions during OGTT. Progesterone administered ip failed to improve glucose excursions during OGTT in WT mice. (Figure 4.11A) To interpret this finding we measured glucose‐stimulated plasma levels of GLP‐1, GIP, insulin and glucagon. Progesterone administered ip significantly increased insulin and GIP levels. In contrast, P4 failed to increase GLP‐1 levels during OGTT (Figure 4.11B) and had no effect on glucagon secretion (Figure 4.11C). Our findings revealed a critical role for enteral progesterone in the regulation of GLP‐1 secretion and glucose homeostasis. Conversely, both enteral and ip progesterone administration increased plasma levels of GIP and insulin during OGTT.

4.5 DISCUSSION To identify molecular mechanisms important for GLP‐1 synthesis and secretion in gut endocrine cells, we and others have studied the genes and proteins expressed in murine GLUTag cells [536]. GLUTag cells retain many of the properties associated with differentiated gut endocrine cells, including cAMP‐dependent and nutrient‐sensitive regulation of proglucagon gene transcription and PGDP secretion [186‐188]. We have now demonstrated that the murine PR, Paqr5 and Paqr7 mRNA transcripts are expressed in different regions of the mouse . Furthermore, progesterone activates ERK1/2 and stimulates GLP‐1 secretion in GLUTag enteroendocrine cells. Moreover, enteral progesterone administration increased GLP‐1 and GIP levels and improved oral glucose tolerance in mice.

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A

0.6 V P4 0.5

0.4

0.3 M x min)M x  ( 0.2

0.1 Acetaminophen (AUC) Acetaminophen 0.0 WT (n=8) DIRKO (n=8)

B 120 120 100 100

80 80 60

60 (%) (%) 40 40 WT -V 20 DIRKO-V 20 WT -P4 DIRKO-P4

Percent fasting glucose 0

Percent fasting glucose fasting Percent 0 0 50 100 150 200 0 50 100 150 200 Time (min) Time (min)

C 4500 V 4000 P4 3500 3000 2500 2000 1500 (mM x min) x (mM AUC glucose 1000 500 0 WT (n=6) DIRKO (n=6)

Figure 4.10 Acute enteral progesterone administration does not modulate the rate of gastric emptying and does not modify insulin sensitivity in mice. (A) AUC for plasma acetaminophen levels in gastric emptying studies with vehicle (V) or progesterone (P4) in WT and DIRKO mice. Liquid‐phase gastric emptying was assessed using the acetaminophen absorption test as described in Methods. (B) Relative decline in glucose levels during an ITT for WT and DIRKO mice pre‐treated with oral vehicle (V) or progesterone (P4) (100 g) 15 min before administration of a single dose of insulin (1.2 U / kg). Values are expressed as a percentage relative to fasting blood glucose levels. (C) AUC for glucose during ITT shown in (B).

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Measure: Insulin and glucagon ip V P4 or Oral glucose -15’ 0’ 5’ 15’ 90’ Time (min) Tail bleed

Measure: Total GLP-1 and Total GIP A D 30 V 50 P4 40 20 30

10 20 Glucose (mM)

tGLP-1 (pg/ml) 10 0 0 20 40 60 80 100 0 Time (min) V P4 B * 1.2 * 800

1.0 600 0.8 400 0.6 tGIP (pg/ml) 200 0.4

Insulin (ng/ml) Insulin 0.2 0 V P4 0.0 V P4 C 120 100 80 60 40

Glucagon (pM) Glucagon 20 0 5 min 15 min

Figure 4.11 Intraperitoneally administered progesterone fails to improve glucose homeostasis during an OGTT in mice. Eight‐ to ten‐week‐old male mice (n=6) were fasted for 4 hours and ip vehicle (dashed line) (day 1) or progesterone (P4) (100 µg) (solid line) (day 2) was administered 15 min before an oral glucose load (OGTT). (A) Glucose excursions during OGTT. Blood was collected at 5 min after glucose administration. This sample was used to measure the plasma levels of (C) Glucagon, D) total GLP‐1 immunoreactivity, (E) total GIP immunoreactivity. A second blood sample was collected at 15 min after glucose administration for the measurement of (B) insulin and (C) glucagon. Vehicle (V) white bars, progesterone (P4) black bars. * p<0.05

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Together, these findings establish a role for progesterone in the control of incretin secretion and glucose homeostasis. Despite expression of a functional PR in GLUTag cells, the ability of R5020 and progesterone to stimulate GLP‐1 secretion was not diminished by the progesterone receptor antagonist RU486. Furthermore, R5020 had no effect on proglucagon promoter‐dependent luciferase activity in GLUTag cells transfected with the ‐ [300]GLU‐Luc plasmid. However, the MAP kinase inhibitor UO126 eliminated the stimulatory effect of progesterone on ERK1/2 phosphorylation and GLP‐1 secretion.

The progesterone‐response element, GGTACAAACTGTTCT, does not appear to be present in the human, mouse and rat proglucagon promoter as assessed by blast analysis (National Center of Biotechnology Information, (NCBI)) of 2500 bases upstream and 100 bases downstream starting from the transcription starting site [537]. However, progesterone has been shown to induce transcription of genes lacking the hormone response element, via alternative genomic and non‐genomic mechanisms, reviewed in [538]. In the present study, we have transfected GLUTag cells with the ‐ [300]GLU‐Luc expression vector, containing the G1, G2, G3, G4 and CREB enhancer elements of the proglucagon promoter. Thus, we cannot discard the possibility that progesterone may stimulate a larger genomic fragment of the proglucagon promoter. However, we demonstrated that P4 did not modulate proglucagon gene expression in mice, and only promoted an increase in proglucagon mRNA transcripts in GLUTag cells when in the presence of FCS. Hence, these results suggest that P4 actions on L‐cells may be restricted to the regulation of GLP‐1 secretion. Together, our findings strongly suggested that progesterone stimulates GLP‐1 secretion through non‐genomic mechanisms linked to ERK1/2 signal transduction. Consistent with this possibility, we observed robust activation of both ERK1/2 phosphorylation and GLP‐1 secretion using BSA‐P4, a hybrid molecule that restricts passage of progesterone across the cell membrane. In contrast, BSA‐P4 had no effect on the transcriptional activity of MMTV‐Luc in GLUTag cells, whereas R5020 robustly increased MMTV‐Luc activity in the same experiments. Limited information is available on whether and how progesterone controls glucose homeostasis or islet function. A previous study demonstrated that female, but not male PR‐/‐ mice exhibited lower fasting glucose and higher insulin levels; however, the precise

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mechanisms accounting for this sexual dimorphism and phenotype were not clearly elucidated [539]. Our data, using both the nuclear progesterone antagonist RU486, and membrane‐bound progesterone, together with siRNA knockdown studies, clearly implicates a role for membrane progesterone receptors in the control of GLP‐1 secretion. The membrane‐restricted actions of progesterone are thought to be linked to the activity of at least three G protein coupled receptors, principally Paqr5, Paqr7, and Paqr8 [438, 540]. Although actions for these receptors have been described in several tissues, principally reproductive organs and the immune system [541], no previous studies have linked membrane progesterone receptors to the control of gut hormone secretion or glucose homeostasis. Intriguingly, although the classical estrogen receptor regulates beta cell function and islet lipid synthesis [542] the membrane estrogen receptor GPR30 also stimulates insulin secretion through non‐genomic pathways in ‐cells [543]. Hence, the mechanisms through which sex steroids regulate glucose homeostasis are complex and include both classical and membrane‐localized receptors. The glucoregulatory actions of enteral P4 were unexpectedly preserved in Glp1r‐/‐, and DIRKO mice; however, they were impaired in Gipr‐/‐ mice. A compensatory up‐regulation of the GIP‐GIPR entero‐insular axis has been described in Glp1r‐/‐ mice and this could explain the preservation of P4‐mediated glucoregulatory actions observed in these mice. Given the widespread distribution of the PR, Paqr5 and Paqr7 expression along the GI tract, it seems likely that progesterone may also influence secretion of other gut‐derived factors like CCK, oxyntomodulin, and PYY that contribute to the control of glucose homeostasis. We could postulate that in DIRKO mice, compensatory mechanisms could enhance the glucoregulatory actions of these gut‐derived factors preserving P4‐mediated glucose control. Furthermore, activation of neuronal circuits cannot be excluded. Intraperitoneal administration of progesterone failed to influence glucose excursions during OGTT despite a significant increase in plasma GIP and insulin levels. Interestingly, ip progesterone failed to increase plasma GLP‐1 levels, hence progesterone‐mediated control of GLP‐1 secretion and possible of other gut peptides, may require direct contact of progesterone with the gut lumen. It is important to consider that in humans oral P4 absorbs poorly, reaches a peak serum level several hours after administration and exhibits great inter‐individual variability [544]. Furthermore, oral‐administered P4 is absorbed in the intestine and it is readily

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metabolized in the liver, hence exhibiting a rapid clearance rate [545]. It is possible that in the experimental design used for these studies, most of the P4 could be expected to remain within the gastrointestinal (GI) tract 15 minutes after oral administration. Hence, P4 may directly stimulate GLP‐1 secretion possibly via activation of the membrane receptors Paqr5 and Paqr7 in the GI tract of mice. If Paqr5 and Paqr7 localized to the apical membrane of enteroendocrine L‐ cells, then enteral P4 could readily activate these receptors and stimulate GLP‐1 secretion in vivo, while ip P4 would fail to do so. Levels of estrogen and progesterone correlate with changes in insulin sensitivity during the menstrual cycle in normal healthy women [546]. Progesterone levels rise during pregnancy, and progesterone is thought to play a role in the development of insulin resistance during the later stages of pregnancy. Indeed direct treatment of adipocytes with progesterone reduced glucose uptake and impaired insulin action in vitro [547]. Analysis of ‐cell function and insulin sensitivity in women treated with progestin‐only contraceptives revealed evidence for reduced insulin sensitivity and compensatory increases in insulin secretion following oral or intravenous glucose administration; however, GLP‐1 levels were not reported in these subjects [548]. Our data expand existing concepts around the control of GLP‐1 secretion and raise interesting questions surrounding a potential role for progesterone in the augmentation of incretin secretion and control of glucose homeostasis under different physiological and pathophysiological situations, including pregnancy and type 2 diabetes.

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Chapter 5

DISCUSSION

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Given that type 2 diabetes is characterized by insulin resistance associated with insufficient β‐cell function and β‐cell mass, pharmacological treatments have been developed to improve insulin sensitivity and to increase insulin secretion. In recent years, therapies that target the GLP‐1R including GLP‐1R mimetics and DPP‐4 inhibitors have been developed [516, 549, 550]. Hence, targeting other insulinotropic GPCRs has received increasing attention, reviewed in [463]. Furthermore, developing incretin secretagogues in combination with DPP‐4 inhibitors could also provide new tools for the treatment of type 2 diabetes. GPR119, an insulinotropic receptor expressed in islet β‐cells and enteroendocrine L and K cells is a target for diabetes drug development. GPR119 actions are not restricted to the islet β‐ cells as its activation promotes incretin secretion from murine enteroendocrine cells in a glucose‐independent manner [341, 344, 515]. However, the precise mechanisms involved in GPR119‐mediated glucose control are not completely understood. Moreover, GPR119 activation was previously shown to reduce the rate of gastric emptying [399]; however, the mechanisms involved were not delineated. In addition, GPR119 activation could directly or indirectly protect islet β‐cells from cytotoxic injury and promote survival. This potential GPR119 action has not been examined in the current literature. In the present work (chapter 2), I have used a combination of pharmacological and physiological experiments to elucidate the mechanisms involved in GPR119‐regulation of glucose homeostasis and modulation of gastric emptying. In chapter 3, using a gain of function method, I assessed the importance of GPR119 signaling for islet β‐cell cytoprotection from STZ‐ induced apoptosis. In addition, using a loss of function model, I evaluated the importance of endogenous GPR119 signaling for β‐cell susceptibility to STZ‐induced apoptosis and for β‐cell function during HF feeding. GLP‐1 and glucagon are derived from the same proglucagon precursor; however, they exert opposite effects on glucose homeostasis [386, 453, 454]. Furthermore, glucagon secretion from islet α‐cells is stimulated in response to hypoglycemia. Conversely, release of GLP‐1 from L‐cells is stimulated by food ingestion. How proglucagon gene transcription and PGDP secretion is differentially regulated in α‐cells and L‐cells is not completely understood. In the present work (chapter 4), using a combination of microarray technology and PCR I have investigated transcription factors differentially expressed in TC1 and enteroendocrine

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GLUTag cells. The result of this study identified PR as a transcription factor expressed in GLUTag cells but not in TC1 cells. Using in vitro methods, I have delineated the signaling pathway involved in P4‐mediated GLP‐1 secretion and I have demonstrated that P4 stimulates GLP‐1 secretion in a glucose‐dependent manner via activation of the membrane PRs: Paqr5 and Paqr7. Furthermore, the work presented in chapter 4 demonstrates that enteral P4 stimulates GLP‐1 and GIP secretion and improves glucose excursions during an OGTT in mice independent of incretin receptor signaling. Below is a brief discussion of the findings presented in this thesis and direction for future work.

What are the mechanisms of GPR119‐dependent control of glucose homeostasis? In chapter 2, I showed that while direct activation of the islet‐GPR119 stimulates insulin secretion in a glucose‐dependent manner in vitro, this action is not sufficient for optimal GPR119‐regulated improvement of glucose control in vivo. Using the DIRKO mouse model I demonstrated that incretin receptor signaling together with enteral glucose exposure is essential for GPR119‐regulated control of glucose homeostasis. However, the insulinotropic actions of GPR119 agonists are independent of incretins as the AR231453‐mediated increase of GSIS is preserved in DIRKO mice. Previous studies suggested that pharmacological activation of GPR119 reduces the rate of gastric empting in rodents [400]. OEA, a natural ligand for GPR119 and PPARα, was shown to reduce the rate of gastric emptying independent of PPARα signaling and its effect was not diminished by co‐administration of the GLP‐1R antagonist exendin (9‐39) [450]. Given that GPR119 activation promotes PGDP secretion from L‐cells and GLP‐1 and GLP‐2 inhibit gastric emptying [461] we hypothesized that GPR119 agonists could modulate gastric emptying by activation of GLP‐1 and/or GLP‐2 signaling. Surprisingly, AR231453 reduced the rate of gastric emptying in WT, Glp1r‐/‐ and DIRKO mice. AR231453 clearly reduced the rate of gastric emptying in Glp2r‐/‐ mice; however, this effect did not reach statistical significance most likely due to the low sample size. PYY, a peptide co‐secreted with GLP‐1 from the enteroendocrine L‐cell is also a downstream target of GPR119 regulation [451]. GPR119 activation was shown to mediate inhibition of gut epithelial electrolyte secretion in a Y1R‐dependent manner [451]. Y1R‐mediated responses are

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predominantly of epithelial nature, while Y2R‐mediated effects are of neuronal origins [505, 551, 552]. In chapter 2 we have clearly demonstrated that AR231453 stimulates PYY secretion in mice during OGTT. In an attempt to determine if PYY signaling was essential for GPR119 transduction of gastric emptying modulation, I co‐administered the Y2R antagonist BIIEO246 with AR231453. Our results demonstrate that AR231453 effects on gastric emptying were not diminished. However, a better model to test the importance of PYY in mediating GPR119 actions on gastric emptying will be to repeat the study using a genetic model of PYY ablation. Taken together these findings demonstrate that many gut enteroendocrine cell peptides known to reduce the rate of gastric emptying like GLP‐1, GLP‐2, and PYY and possible other yet unidentified peptides are downstream targets of GPR119 signaling. Hence, elimination of a single peptide does not abrogate AR231453 inhibition of gastric emptying suggesting the involvement of multifactorial mechanisms. In conclusion, the results presented in chapter 2 of this thesis demonstrate that GPR119 agonists regulate glucose via complementary mechanisms: direct β‐cell insulinotropic actions, incretin secretagogue actions and reduction of the rate of gastric emptying.

Does GPR119 activation exert β‐cell cytoprotective actions in vivo? GLP‐1 and GIP both stimulate cAMP production. In the β‐cell GLP‐1 and to a lesser extent GIP have been shown to exert cytoprotective actions [118, 119, 132, 133, 396]. Furthermore, abrogation of endogenous GLP‐1 signaling results in reduced number of large islets and increased susceptibility to β‐cell apoptosis [132, 492]. GPR119 also couples to Gαs and stimulates cAMP accumulation; hence, it could potentially play a role in β‐cell survival. Furthermore, GPR119 agonists stimulate GLP‐1 secretion [340, 341, 344, 515]; hence, activation of GPR119 could indirectly play a role in β‐cell cytoprotection. Moreover, mice lacking GPR119 exhibit reduced GLP‐1 levels [387]. Hence, elimination of endogenous GPR119 signaling could result in enhanced susceptibility to β‐cell apoptosis via direct or indirect mechanisms. In chapter 3, I addressed these questions. Using the small synthetic molecule, specific agonist for GPR119, AR881, I demonstrated that sustained administration of AR881 (8 days) did not exert cytoprotective actions in the low dose STZ model of β‐cell injury. A recent study done in rat insulinoma‐derived β‐cell lines BRIN‐BD11 and INS‐1E [553, 554] exposed to palmitate, showed that OEA exerted cytoprotective actions independent of GPR119 signaling [555].

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The authors identified the presence of GPR119 transcripts in both cell lines; however, they were not able to demonstrate that the receptor was functional. Furthermore, specific GPR119 agonists like AR231453 failed to protect these cells from the deleterious actions of palmitate. In my study, I have not assessed circulating incretin levels following AR881 treatment, which could have given an indication of compound efficacy. However, I tested the effect of AR881 by acute administration during OGTT and demonstrated that AR881 controlled glucose homeostasis. Nevertheless, it is possible that in our experimental design we could have induced GPR119 desensitization. The natural ligand OEA and the specific synthetic ligands PSN632408 and PSN375963 all showed bell‐shape dose response profiles in vitro [337, 341, 505] indicating the possibility of GPR119 desensitization. Furthermore, it is possible that our experimental protocol could have resulted in reduced GPR119 mRNA transcripts or protein levels. To investigate this possibility it will be worthwhile to repeat the present study using at least two independent GPR119 specific agonists. The experimental end‐points, apart from the assessment of β‐cell apoptosis, should include measurement of incretin levels and collection of intestine and pancreatic tissue for the assessment of GPR119 expression.

What is the role of endogenous GPR119 signaling for β‐cell biology? It has been reported that islets from Glp1r‐/‐ mice exhibit reduced mRNA transcript levels of Igf1r, IRS2 and EGFR compared to WT controls suggesting that GLP‐1R may modulate the sensitivity of islet β‐cells to cytoprotective pathways [396, 506]. In the present work (chapter 3), I examined whether the absence of physiological GPR119 signaling modifies the mRNA transcript levels of selected genes important for β‐cell biology. I demonstrated that under basal conditions, islets isolated from young (7 weeks old) GPR119‐/‐ mice, exhibit reduced mRNA levels of Irs2, Glp1r, and Vipr1. These findings suggest that endogenous GPR119 signaling may modulate β‐cell cytoprotective pathways. To determine the mechanisms involved (direct vs. indirect) a comparative study using conditional β‐cell specific vs L‐cell specific GPR119 knockout mouse is warranted. Furthermore, introducing GPR119 back into islets isolated from a conditional β‐cell specific GPR119 knockout mouse will also shed light into the importance of GPR119 signaling in the maintenance of basal expression of key genes important for β‐cell function and survival independent of possible compensatory mechanisms that could play a role in islets from the global GPR119 knockout mouse.

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In chapter 3, I demonstrated that endogenous GPR119 signaling influences the susceptibility of islet β‐cells to STZ‐induced apoptosis and to the deleterious actions of HF feeding. I reported that β‐cells from GPR119‐/‐ mice exhibited enhanced sensitivity to STZ‐induced apoptosis compared to WT mice. Furthermore, GPR119‐/‐ mice displayed impaired β‐cell function when fed a HFD compared to WT controls. Moreover, I have shown that islets from GPR119‐/‐ mice fed a HFD failed to increase mRNA levels of selected genes known to be important for β‐cell survival and function compared to WT controls. Hence, it is possible that lack of GPR119 signaling results in a β‐cell nutrient sensing defect. I have not detected significant differences in pancreatic insulin levels nor in islet size and number in GPR119‐/‐ compared to WT controls under RC and HF feeding conditions. However, since I have not measured pancreatic weight, I have not been able to determine β‐cell mass in the present work. Moreover, a limitation of the described experiment is that WT mice fed a HF diet did not display a significant increase in β‐cell area and islet number, nor in insulin content and they did not show significant impairment in glucose control compared to RC fed mice. A possible explanation resides in the RC diet used in this study. We used standard RC provided by our animal housing facility. This diet contains 18% Kcal from fat sources. Consequently, mice become heavier than expected masking the impact of the chosen HFD (45% Kcal from fat) on the parameters measured in this study. We are currently repeating this study using a RC (10 % Kcal from fat) versus HFD (45 % Kcal from fat). Nevertheless, the results presented in chapter 3 suggest that GPR119 signaling is not essential for insulin synthesis, islet size and β‐cell adaptation to HF feeding. I have shown that islets from GPR119‐/‐ mice exhibit reduced Gcg mRNA levels compared to WT islets. However, in these studies, I have not investigated the number, area and localization of islet α‐cells. Hence, a detail assessment of islet α‐cell topography may be useful in interpreting this finding. Furthermore, assessment of Gcg mRNA levels in islets isolated from older mice did not display differential transcript levels. This could be due to the low number of mice used in this study (n=3‐4). In addition to investigating mRNA levels, a more detailed study assessing protein levels of glucagon, intra‐islet GLP‐1 and PC1/2 would be useful in evaluating possible compensatory mechanisms during aging in islets of GPR119‐/‐ mice.

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OEA, PSN632408 and AR231453 stimulate β‐cell proliferation and improve islet graft function in mice [280, 556]. However, in these studies, GLP‐1 levels were significantly higher in mice treated with the GPR119 agonists compared to controls. Hence, the improvement in islet graft could have been due to enhanced GLP‐1 signaling. Furthermore, there is no current evidence showing a direct action of GPR119 agonists on β‐cell proliferation under basal conditions and further investigation is required. In summary, our results indicate a critical role, not previously identified, for endogenous GPR119, directly or indirectly, protecting β‐cells from STZ‐induced apoptosis and maintaining adequate β‐cell function during HF feeding.

Novel progesterone mediated pathway for the control of GLP‐1 secretion The work presented in chapter 4 provides evidence for a novel role for P4 via non‐genomic actions in the regulation of GLP‐1 secretion. I have demonstrated that P4 stimulates GLP‐1 secretion in GLUTag cells. The signal transduction pathway involves activation of ERK1/2 and is dependent on the membrane receptors Paqr5 and Paqr7. Furthermore, P4‐mediated GLP‐1 secretion is independent of the PR antagonist RU486 both in vitro and in vivo, suggesting that PR is not involved in these actions. It is important to mention that we have not assessed the efficacy of oral RU486 to inhibit PR in our in vivo studies. I have shown that in male mice, P4 stimulates GLP‐1, GIP and insulin secretion resulting in reduced glucose excursions during OGTT. Unexpectedly, P4‐mediated glucose control was preserved in Glp1r‐/‐ and DIRKO mice while it was lost in Gipr‐/‐ mice. Hence, other gut peptides or factors may be stimulated by enteral P4 and be involved in the glucoregulatory actions seen in mice. Moreover, it will be important to follow up these studies assessing levels of glucagon following P4 administration during OGTT in Gipr‐/‐ mice. Interestingly, enteral administration of P4 is essential for glucose control, since ip P4 failed to improve glucose homeostasis. Furthermore, while ip P4 increased GIP levels, the levels of GLP‐1 remained unchanged, suggesting the need for enteral P4 for the regulation of peptide secretion from the enteroendocrine L‐cell. It is possible that P4 stimulates GLP‐1 secretion directly from the enteroendocrine L cell in mice via activation of the membrane‐bound receptors Paqr5 and Paqr7, as is the case with GLUTag cells and if these receptors localize to the apical membrane of L‐cells, then, lumen exposure to P4 could be required for P4‐mediated

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stimulation of GLP‐1 secretion. Finally, in the results described in chapter 4, I have presented evidence that demonstrates that acute P4 does not reduce the rate of gastric emptying and does not impair insulin sensitivity. In pregnancy, the levels of P4 increase to ensure embryo implantation [557]. In mammals, including humans, pregnancy promotes dramatic maternal metabolic changes including increased glucose‐stimulated insulin secretion [558]. In addition, pregnancy is associated with an increase in hepatic glucose production to ensure proper nutrient fetal supply. However, the last trimester is characterized by the development of insulin resistance that in some individuals progresses to gestational diabetes [559]. The development of insulin resistance and gestational diabetes is believed to be associated with the elevated circulating levels of P4. For example, studies in ovariectomized rats treated with P4 demonstrated reduced muscle‐ glucose uptake, reduced GLUT4 protein expression and increased insulin resistance during exercise [560]. In addition, adipocytes isolated from ovariectomized rats, chronically treated with P4, exhibit a reduced rate of glucose oxidation and increased insulin resistance [561, 562]. Moreover, chronic P4 administration to db/db female mice resulted in a significant increase in fasting glucose levels compared to vehicle‐treated mice [539]. Interestingly, in the same study, administration of the PR antagonist RU486 decreased fasting glucose levels in WT and db/db female mice. Furthermore, fasting glucose levels were lower in female mice lacking the PR compared to WT controls. In addition, PR‐/‐ females had improved glucose tolerance and increased glucose‐stimulated insulin during IPGTT [539]. Hematoxylin/eosin staining and double immunofluorescent staining of pancreatic sections with antibodies, directed against glucagon and insulin, demonstrated that islets from PR‐/‐ female mice were larger and contained increased number of β‐cells, but no changes in the number of α‐cells compared to PR+/+ islets.

In the latter study, the levels of P4 and gut hormones (GLP‐1, GIP) were not assessed. I have shown that P4 stimulates GLP‐1 secretion independent of PR and via Paqr5 and Paqr7 activation. Hence, we could hypothesize that in the absence of PR signaling (like in the PR‐/‐ mouse model); the desirable actions of P4 could be unmasked. A future study could examine whether the improved glucose control seen in PR‐/‐ females is the result of enhanced incretin secretion.

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There are not many studies investigating the correlation between progesterone levels and gut hormones. A study done in pregnant rats has found a correlation between P4 and PYY levels throughout pregnancy [563]. Moreover, a study in humans has found that the levels of PYY secreted during OGTT were significantly higher in females than males [467]. Since PYY is co‐secreted with GLP‐1 [467], it is expected that a similar correlation exists between P4 levels and GLP‐1 levels in pregnancy. Another important finding is the observation that only enteral and not ip P4 stimulates GLP‐1 secretion in vivo. This is of importance, especially for women undergoing hormone replacement therapy. A future study in humans could evaluate incretin levels and their correlation to P4 levels during OGTT in this population. Overall, my data confirmed that GPR119 activation stimulates incretin secretion, and has revealed that GPR119 agonists require intact incretin signaling for optimal glucose control in vivo. My studies have also confirmed that GPR119 agonists inhibit the rate of gastric emptying in a GPR119‐dependent, yet incretin, PY2R‐ and GLP‐2R‐independent manner. I have demonstrated that mice lacking endogenous GPR119 signaling exhibit enhanced islet β‐cell susceptibility to STZ‐induced apoptosis. My studies using GPR119‐/‐ mice fed a HFD have revealed a potential role, previously unidentified, for GPR119 signaling in β‐cell adaptation to nutrient excess. I have provided, for the first time, a detailed analysis of GPR119‐/‐ islet mRNA transcripts compared to GPR119+/+ islets, under basal and HFD conditions. I believe that my results will set the foundation for future studies needed to further elucidate the role of GPR119 in islet β‐cell function, proliferation and survival. Taken together, these studies are useful contributions to our understanding of GPR119 biology. Understanding GPR119 biology is of particular importance since GPR119 represent a target to improve glucose homeostasis in type 2 diabetic subjects due to the combined stimulation of insulin and incretin secretion observed with GPR119 agonists. Thus, many pharmaceutical companies are pursuing GPR119 agonists as potential therapeutic tools for the treatment of Type 2 diabetes, for a review see [391, 564]. Phase II clinical trials of MBX2908 reported improved glucose control and stimulated GLP‐1 levels in normoglycemic individuals. In individuals with increased fasting glucose, MBX2982 improved glucose control following a meal and enhanced insulin release following glucose infusions [565]. Phase I clinical trials of APD‐597 provided evidence for improvement of

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post‐meal glucose excursions and stimulation of GLP‐1, GIP and PYY in both overweight and obese non‐diabetic volunteers and in subjects with type 2 diabetes [566], for a review see [567] In this thesis I have also presented a novel pathway involving the activation of Paqr5 and Paqr7 that suggests a previously unsuspected role for progesterone in the regulation of GLP‐1 secretion and glucose homeostasis. These results raise the possibility for intestinal membrane progesterone receptors as novel targets for the development of selective agonists that augment incretin secretion and control glucose homeostasis, independent of systemic progesterone exposure. These findings could have implications in physiological and pathophysiological conditions such as pregnancy, menopause, and gestational diabetes.

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