Pancreatic Beta Cell in Health and Disease

Susumu Seino, Graeme I. Bell (Eds.) Pancreatic Beta Cell in Health and Disease Susumu Seino, Graeme I. Bell (Eds.) P...

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Susumu Seino, Graeme I. Bell (Eds.) Pancreatic Beta Cell in Health and Disease

Susumu Seino, Graeme I. Bell (Eds.)

Pancreatic Beta Cell in Health and Disease

Susumu Seino, M.D., D.M. Sci. Professor, Division of Cellular and Molecular Medicine, Department of Physiology and Cell Biology, Kobe University Graduate School of Medicine 7-5-1 Kusunoki-cho, Chuo-ku, Kobe 650-0017, Japan Graeme I. Bell, Ph.D. Louis Block Distinguished Service Professor, Medicine and Human Genetics, The University of Chicago 5841 South Maryland Avenue, Chicago, IL 60637, USA

ISBN 978-4-431-75451-0

e-ISBN 978-4-431-75452-7

Library of Congress Control Number: 2007939279 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, speciﬁcally the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microﬁlms or in other ways, and storage in data banks. The use of registered names, trademarks, etc. in this publication does not imply, even in the absence of a speciﬁc statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Product liability: The publisher can give no guarantee for information about drug dosage and application thereof contained in this book. In every individual case the respective user must check its accuracy by consulting other pharmaceutical literature. Springer is a part of Springer Science+Business Media springer.com © Springer 2008 Printed in Japan Typesetting: SNP Best-set Typesetter Ltd., Hong Kong Printing and binding: Shinano Co. Ltd., Japan Printed on acid-free paper

Preface

The beta cells of the pancreatic islets of Langerhans are the only cells in the body that produce and secrete insulin, a key metabolic hormone, which plays a central role in the maintenance of glucose homeostasis regulating glucose uptake in muscle and adipose tissue as well as carbohydrate, fat, and protein metabolism in these tissues and the liver. Failure of normal beta-cell function can lead to hyperglycemia (diabetes mellitus) or hypoglycemia. While hypoglycemia is a rare and life-threatening condition in which the beta cell secretes too much insulin, diabetes mellitus is a common disorder that is fast becoming an epidemic. It is a major threat to human health in the 21st century. The number of diabetic patients worldwide is rapidly increasing and is predicted to reach 380 million by 2025, according to the International Diabetes Federation (IDF). Diabetes mellitus is a heterogeneous disorder with some forms such as maturity-onset diabetes of the young and permanent neonatal diabetes being primary genetic disorders of the beta cell. Monogenic forms of diabetes are an uncommon cause of diabetes representing about 1% of cases. The common forms of diabetes mellitus, designated type 1 and type 2, are multifactorial in origin with both genetic and environmental factors contributing to their development. Type 1 diabetes is caused by autoimmune destruction of beta cells leading to an absolute deﬁciency of insulin and fatal hyperglycemia and ketoacidosis if not treated. Type 2 diabetes is a disorder of relative deﬁciency of insulin resulting when the beta-cell mass is not able to expand and thereby secrete more insulin in response to an increasing demand such as occurs in obese individuals. Whatever the form of diabetes, the beta cell plays a central role in the disease process. The purpose of this book is to provide a comprehensive up-to-date review of the beta cell in health and disease. The chapters address the architecture and pathology of normal and diabetic pancreatic islets; regulation of beta-cell proliferation and death; the potential of stem cells and extra-pancreatic

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Preface

tissues as beta-cell replacement therapies; insulin biosynthesis from transcription to processing; regulation of insulin secretion including metabolic control, ion channels, gap junctions, cyclic AMP, incretins and exocytosis; development of novel imaging techniques to visualize the exocytosis of insulin granules; and genetic disorders of the beta cell. We hope this book inspires students and young basic and clinical investigators to become actively involved in diabetes research and the search for new approaches for preventing and treating diabetes. In addition, we hope it will provide encouragement to others in the ﬁeld. Although we have made a great deal of progress in understanding the relationship between the beta cell and health and disease, much remains to be done. We gratefully acknowledge the contribution to this book by each of the authors. We thank the editorial staff of Springer Japan who patiently assisted us throughout the project. Without their effort, this book would not have been possible.

Editors

Susumu Seino, M.D., D.M. Sci. Division of Cellular and Molecular Medicine Department of Physiology and Cell Biology Kobe University Graduate School of Medicine Kobe, Japan

Graeme I. Bell, Ph.D. Departments of Medicine and Human Genetics The University of Chicago Chicago, IL, USA

Contents

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I: Pancreatic Beta Cell and Insulin Biosynthesis 1. Architecture of Pancreatic Islets M. Brissova and A.C. Powers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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2. Transcriptional Regulation of Insulin Gene Expression I. Artner and R. Stein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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3. The Biosynthesis of Insulin D.F. Steiner . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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II: Cell Signaling and Insulin Secretion 4. Metabolic Regulation of Insulin Secretion B.E. Corkey . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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5. Mechanisms of Beta-Cell Death in Diabetes M.Y. Donath and J.A. Ehses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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6. Ion Channels and Insulin Secretion D.A. Jacobson and L.H. Philipson . . . . . . . . . . . . . . . . . . . . . . . . . . .

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7. Gap Junctions and Insulin Secretion P. Klee, S. Bavamian, A. Charollais, D. Caille, J. Cancela, M. Peyrou, and P. Meda . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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8. Protein Kinase A-Independent Mechanism of cAMP in Insulin Secretion S. Seino, T. Miki, and T. Shibasaki . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contents

9. Regulation of Insulin Granule Exocytosis E. Renström and P. Rorsman . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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10. Mechanism of Insulin Exocytosis Analyzed by Imaging Techniques S. Nagamatsu and M. Ohara-Imaizumi . . . . . . . . . . . . . . . . . . . . . . .

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11. Two-Photon Excitation Imaging of Insulin Exocytosis N. Takahashi and H. Kasai . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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III: Pancreatic Development and Beta-Cell Formation 12. Regulation of Beta-Cell Growth and Death C.J. Rhodes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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13. Beta-Cell Replication S.J. Salpeter and Y. Dor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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14. Stem Cells as a Cure for Diabetes T. Otonkoski, M. Banerjee, and K. Lundin . . . . . . . . . . . . . . . . . . . .

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15. Use of Extra-Pancreatic Tissues for Cell Replacement Therapy for Diabetes I. Meivar-Levy and S. Ferber . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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IV: Incretins and Beta-Cell Function 16. Molecular Biology of Gluco-Incretin Function S. Klinger and B. Thorens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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17. Incretins and Regulation of Insulin Secretion M.A. Nauck and J.J. Meier . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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V: Pancreatic Beta Cell and Disease 18. Pancreatic Islet Pathology in Type 2 Diabetes A. Clark . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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19. Genetic Disorders of the Pancreatic Beta Cell and Diabetes (Permanent Neonatal Diabetes and Maturity-Onset Diabetes of the Young) E.L. Edghill and A.T. Hattersley . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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20. ATP-Sensitive Potassium Channels in Health and Disease P. Proks and F.M. Ashcroft . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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21. Glucokinase in Glucose Homeostasis, Diabetes Mellitus, Hypoglycemia, and as Drug Receptor F.M. Matschinsky . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors

Artner, Isabella (p. 13) Department of Molecular Physiology and Biophysics, Vanderbilt University Medical Center, 723 Light Hall, Nashville, TN 37232, USA Ashcroft, Frances M. (p. 431) Oxford Centre for Gene Function, Department of Physiology, University of Oxford, Parks Road, Oxford OX1 3PT, UK Banerjee, Meenal (p. 265) Hospital for Children and Adolescents and the Biomedicum Stem Cell Center, Biomedicum Helsinki, P.O. Box 63, Haartmaninkatu 8, FI-00014 University of Helsinki, Helsinki, Finland Bavamian, Sabine (p. 111) Department of Cell Physiology and Metabolism, University of Geneva, Medical School, 1 rue Michel Servet, 1211 Genève 4, Switzerland Brissova, Marcela (p. 3) Division of Diabetes, Endocrinology, and Metabolism, Department of Medicine, Vanderbilt University Medical Center, Nashville, TN, USA Caille, Dorothée (p. 111) Department of Cell Physiology and Metabolism, University of Geneva, Medical School, 1 rue Michel Servet, 1211 Genève 4, Switzerland Cancela, José (p. 111) Department of Cell Physiology and Metabolism, University of Geneva, Medical School, 1 rue Michel Servet, 1211 Genève 4, Switzerland

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Charollais, Anne (p. 111) Department of Cell Physiology and Metabolism, University of Geneva, Medical School, 1 rue Michel Servet, 1211 Genève 4, Switzerland Clark, Anne (p. 381) Diabetes Research Laboratories, Oxford Centre for Diabetes, Endocrinology and Metabolism, Churchill Hospital, Oxford OX3 7LJ, UK Corkey, Barbara E. (p. 53) Obesity Research Center, Department of Medicine, Boston University School of Medicine, Boston, MA 02118, USA Donath, Marc Y. (p. 75) Clinic of Endocrinology and Diabetes, University Hospital Zurich, CH-8091 Zurich, Switzerland Dor, Yuval (p. 245) Department of Cellular Biochemistry and Human Genetics, The Hebrew University–Hadassah Medical School, Jerusalem 91120, Israel Edghill, Emma L. (p. 399) Institute of Biomedical and Clinical Science, Peninsula Medical School, Barrack Road, Exeter, Devon EX2 5AX, UK Ehses, Jan A. (p. 75) Clinic of Endocrinology and Diabetes, University Hospital Zurich, CH-8091 Zurich, Switzerland Ferber, Sarah (p. 285) Endocrine Institute, Sheba Medical Center, Tel-Hashomer 52621, Israel Department of Human Genetics and Molecular Medicine, Sackler School of Medicine, Tel-Aviv University, Tel-Aviv, Israel Hattersley, Andrew T. (p. 399) Institute of Biomedical and Clinical Science, Peninsula Medical School, Barrack Road, Exeter, Devon EX2 5AX, UK Jacobson, David A. (p. 91) Department of Medicine, The University of Chicago, 5841 S. Maryland Ave., MC1027, Chicago, IL 60637, USA

Contributors

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Kasai, Haruo (p. 195) Division of Biophysics, Center for Disease Biology and Integrative Medicine, Faculty of Medicine, University of Tokyo, 7-3-1 Hongo, Bunkyoku, Tokyo 113-0033, Japan Klee, Philippe (p. 111) Department of Cell Physiology and Metabolism, University of Geneva, Medical School, 1 rue Michel Servet, 1211 Genève 4, Switzerland Klinger, Sonia (p. 315) Department of Physiology and Center for Integrative Genomics, University of Lausanne, Genopode Building, Lausanne, Switzerland Lundin, Karolina (p. 265) Hospital for Children and Adolescents and the Biomedicum Stem Cell Center, Biomedicum Helsinki, P.O. Box 63, Haartmaninkatu 8, FI-00014 University of Helsinki, Helsinki, Finland Matschinsky, Franz M. (p. 451) Department of Biochemistry and Biophysics, University of Pennsylvania, School of Medicine, 501 Stemmler Hall, 36th & Hamilton Walk, Philadelphia, PA 19104, USA Meda, Paolo (p. 111) Department of Cell Physiology and Metabolism, University of Geneva, Medical School, 1 rue Michel Servet, 1211 Genève 4, Switzerland Meier, Juris J. (p. 335) Medizinische Klinik I, St. Josef-Hospital, Klinikum der Ruhr-Universität Bochum, Gudrunstrasse 59, D-44791 Bochum, Germany Meivar-Levy, Irit (p. 285) Endocrine Institute, Sheba Medical Center, Tel-Hashomer 52621, Israel Miki, Takashi (p. 133) Division of Cellular and Molecular Medicine, Kobe University Graduate School of Medicine, Kobe 650-0017, Japan Nagamatsu, Shinya (p. 177) Department of Biochemistry, Kyorin University School of Medicine, 6-20-2 Shinkawa, Mitaka, Tokyo 181-8611, Japan

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Contributors

Nauck, Michael A. (p. 335) Diabeteszentrum Bad Lauterberg, Kirchberg 21, D-37431 Bad Lauterberg, Germany Ohara-Imaizumi, Mica (p. 177) Department of Biochemistry, Kyorin University School of Medicine, 6-20-2 Shinkawa, Mitaka, Tokyo 181-8611, Japan Otonkoski, Timo (p. 265) Hospital for Children and Adolescents and the Biomedicum Stem Cell Center, Biomedicum Helsinki, P.O. Box 63, Haartmaninkatu 8, FI-00014 University of Helsinki, Helsinki, Finland Peyrou, Manon (p. 111) Department of Cell Physiology and Metabolism, University of Geneva, Medical School, 1 rue Michel Servet, 1211 Genève 4, Switzerland Philipson, Louis H. (p. 91) Department of Medicine, The University of Chicago, 5841 S. Maryland Ave., MC1027, Chicago, IL 60637, USA Powers, Alvin C. (p. 3) Division of Diabetes, Endocrinology, and Metabolism, Department of Medicine, Department of Molecular Physiology and Biophysics, Vanderbilt University Medical Center, Nashville, TN, USA VA Tennessee Valley Healthcare System, Nashville, TN, USA Proks, Peter (p. 431) Oxford Centre for Gene Function, Department of Physiology, University of Oxford, Parks Road, Oxford OX1 3PT, UK Renström, Erik (p. 147) Lund University Diabetes Centre/Department of Clinical Sciences, UMAS Entrance 72, CRC 91-11, SE20502 Malmö, Sweden Rhodes, Christopher J. (p. 215) Department of Medicine, Section of Endocrinology, Diabetes, and Metabolism, University of Chicago, 5841 S. Maryland Avenue, MC 1027, Room N138, Chicago, IL 60637, USA

Contributors

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Rorsman, Patrik (p. 147) Oxford Centre for Diabetes, Endocrinology and Metabolism, University of Oxford, Oxford OX3 7LJ, UK Salpeter, Seth J. (p. 245) Department of Cellular Biochemistry and Human Genetics, The Hebrew University–Hadassah Medical School, Jerusalem 91120, Israel Seino, Susumu (p. 133) Division of Cellular and Molecular Medicine, Kobe University Graduate School of Medicine, Kobe 650-0017, Japan Shibasaki, Tadao (p. 133) Division of Cellular and Molecular Medicine, Kobe University Graduate School of Medicine, Kobe 650-0017, Japan Stein, Roland (p. 13) Department of Molecular Physiology and Biophysics, Vanderbilt University Medical Center, 723 Light Hall, Nashville, TN 37232, USA Steiner, Donald F. (p. 31) Department of Biochemistry and Molecular Biology, The University of Chicago, 5841 S. Maryland Avenue, MC 1027, Rm N216, Chicago, IL 60637, USA Takahashi, Noriko (p. 195) Division of Biophysics, Center for Disease Biology and Integrative Medicine, Faculty of Medicine, University of Tokyo, 7-3-1 Hongo, Bunkyoku, Tokyo 113-0033, Japan Thorens, Bernard (p. 315) Department of Physiology and Center for Integrative Genomics, University of Lausanne, Genopode Building, Lausanne, Switzerland

I. Pancreatic Beta Cell and Insulin Biosynthesis

1. Architecture of Pancreatic Islets Marcela Brissova* and Alvin C. Powers*,†,‡

Summary. Pancreatic islets are highly vascularized mini-organs that include endocrine cells, endothelial cells, and extracellular matrix. The three-dimensional islet structure and relationship between endocrine cells are different in rodent islets and human islets. This chapter reviews recent studies on the morphology, cell composition, and cell arrangement in human islets and how this may affect islet function.

Introduction The islets of Langerhans or pancreatic islets are scattered throughout the pancreas and constitute about 1%–2% of the pancreas. Islets contain several different cell types—including endocrine cells, endothelial cells, nerves, and ﬁbroblasts—and range in size from just a few islet cells to a complex of several thousand islet cells with a diameter up to 300–400 µm [1]. An incompletely deﬁned and variable capsule encloses an islet and partially separates endocrine cells from exocrine cells. While most islets are <160 µm in diameter, islets >160 µm in diameter are responsible for the majority of total islet mass or volume [1]. Pancreatic islets are highly vascularized mini organs (Fig. 1) that receive a disproportionately larger fraction of pancreatic blood ﬂow (up to 5%–10% of total pancreatic blood ﬂow) than the exocrine portion of the pancreas [2,3]. This chapter focuses primarily on recent work on the endocrine cell composition and architecture of human islets with an emphasis on studies using confocal microscopy.

*Division of Diabetes, Endocrinology, and Metabolism, Department of Medicine, †Department of Molecular Physiology and Biophysics, Vanderbilt University Medical Center, Nashville, TN, USA ‡ VA Tennessee Valley Healthcare System, Nashville, TN, USA

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Fig. 1A,B. Islets of Langerhans—highly vascularized mini-organs. Three-dimensional reconstruction of optical sections through the mouse pancreas. A Sections labeled for insulin (green), glucagon (blue), and vascular endothelial cell marker CD31 (red). B Islet vasculature revealed in more detail by endothelium-binding lectin-FITC (green). Reproduced from [27]

Pancreatic Islet Cell Composition, Architecture, and Arrangement Pancreatic islets were classically described to contain four different cell types: insulin-producing beta cells, glucagon-producing alpha cells, somatostatinproducing delta cells, and pancreatic polypeptide-producing PP cells [1]. Recently, a ﬁfth islet cell type, ghrelin-producing cells, was discovered and termed epsilon cells [4–7]. In both mouse and human, numerous epsilon cells are present in islets during pancreas development and at birth. However, it appears that their population declines postnatally [5,8]. Several approaches, including immunocytochemistry for islet hormones and electron microscopic examination of the distinctive secretory granules for each islet hormone, have been used to study the endocrine cell composition and arrangement of islets. Both approaches require physical sectioning of the pancreas, and careful sampling and reconstruction to avoid bias because of several confounding issues. For example, histologic sections for light microscopy are 5–10 µm in thickness and electron microscopy (EM) sections are even thinner. Thus, the ability to completely examine all sections that would span a single islet by such techniques is laborious and quite difﬁcult. Furthermore, one cannot be certain if a section being examined is through the “top,” “middle,” or “bottom” of an islet. The peripheral location of non-beta cells may give the appearance that non-beta cells are in the “middle” of an islet. Finally, there may be variations in the islet cell composi-

Architecture of Pancreatic Islets

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tion in different regions of the pancreas (discussed below) and thus, examination of multiple sections from different pancreatic regions is required. Using these approaches, the arrangement of endocrine cell types in mammalian islets, originating from studies in primarily rodents, has been referred to as core–mantle architecture with beta cells located centrally and non-beta islet cells being around the islet periphery [1,3,9,10]. A similar histologic approach to study human pancreata, primarily from autopsy specimens, suggested that human islets differed in cell arrangement from the more widely studied rodent islet [11,12]. The physical relationship between the different endocrine cell types has physiologic signiﬁcance since endocrine cells likely communicate by cell–cell interactions, by paracrine mechanisms, and by secretion of hormones and products from one endocrine cell type inﬂuencing “downstream” endocrine cells. Two recent developments have provided the opportunity to investigate human pancreatic islet morphology in more detail. First, the increased availability of human pancreas specimens and isolated human islets for transplantation and research allowed investigators to reassess human islet architecture more rigorously and in larger number of pancreata. Secondly, laser scanning confocal microscopy was utilized to determine unambiguously the arrangement of beta and non-beta cells within islets of different species with an emphasis on human islets [13,14]. In contrast to conventional ﬂuorescence microscopy, laser scanning confocal micro-scopy acquires optical sections through the specimen and thus overcomes the challenge of creating and examining a large number of serial histological sections. By applying this approach to both pancreatic specimens and isolated islets, investigators have described striking interspecies differences in islet architecture (Fig. 2) [13,14]. In sharp contrast to mouse, human islets appear to be quite heterogeneous in their composition as illustrated by data in Fig. 3A and islet images in Fig. 3B and C. While normal adult murine islets have a beta cell core and non-beta cell mantle, human islets do not have the core– mantle architecture and beta cells are rather intermingled with other endocrine cell types, as demonstrated by co-localization for beta cell-produced insulin, alpha cell-produced glucagon and delta cell-produced somatostatin. A similar appearance had been noted by Brelje and colleagues with an early generation confocal microscope, but this examination was limited by the depth of penetration within the islet [15]. Interestingly, porcine, canine, and non-human primate islets [13,14,16] display architecture similar to human islets. Other investigators have suggested that, even though the histologic appearance of human islets is different, the arrangement of endocrine cells in human islets is not random or intermingled, but is in functional units within an islet and that units maintain the core–mantle relationship seen in rodent islets [1,3].

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B

Fig. 2A,B. Interspecies differences in islet architecture. In contrast to mouse, beta cells are intermingled with alpha and delta cells in non-human primate and human islets. A Series of optical sections through the entire isolated mouse (top panel), non-human primate (rhesus macaques) (middle panel), and human islet (bottom panel) acquired at 1-µm intervals in axial (z) dimension (only small subset of optical sections is shown). The stack of optical sections was 3-D reconstructed and the islet projection is shown on far right of each panel. Beta cells (green); alpha cells (red); delta cells (blue). Reproduced from [13]. B Confocal micrographs of islets in the human (upper left), non-human primate (cynomolgus monkey) (upper right), mouse (lower left), and porcine (lower right) pancreas. Beta cells (red); alpha cells (green); delta cells (blue). Reproduced from [14]

Architecture of Pancreatic Islets

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Fig. 3A–E. Adult human islets have fewer beta cells and are heterogeneous in composition compared to mouse islets. A Morphometric analysis of cell composition with respect to beta, alpha and delta cells in the isolated mouse and human islets. B, C Example of two isolated human islets with different cell composition. Beta cells (green); alpha cells (red); delta cells (blue). A–C Reproduced from [13]. D Islet composition with respect to insulin-, glucagon-, and somatostatin-producing cells (beta, alpha, and delta cells, respectively) measured in the human pancreatic sections acquired from four different regions of the human pancreas. E Comparison of islet cell composition derived from morphometric analysis of human and mouse pancreatic sections. D, E Reproduced from [14]

Adult Human Islets Have Fewer Beta Cells, but More Alpha Cells, than Mouse Islets Human islets not only have an architecture distinct from that of rodent islets, but endocrine cell populations are also quite different in human islets (Fig. 3). This was determined by morphometric examination of either

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whole-mount islets (Fig. 3A) [13] or pancreatic specimens (Fig. 3D and E) [14] subjected to laser scanning confocal microscopy. In contrast to prior literature [17–19], these studies indicated that the relative proportion of beta cells in the human islets is much less than in murine islets (approximately 55% versus 77%). On the other hand, the fraction of alpha cells is greater in human islets than in murine islets (approximately 35% versus 18%). The lower fraction of beta cells and more alpha cells in adult human islets were also found by ﬂow cytometry of dispersed islets used for human islet transplantation [20,21] and by morphometry on histological sections of human pancreas [22]. Rat and mouse islets have a homogeneous and constant beta cell distribution [13,14,23] in contrast to wide variation in beta and alpha cell percentage found within individual human and canine islets [13,14,16] and described in PP-rich and PP-poor regions of the human and canine pancreas [16,24,25]. These recent studies of human islets are further strengthened because multiple islet preparations from different islet isolation centers were studied [13,14]. Also, since these islets were isolated from the whole pancreas, they are likely representative of islet population within the pancreas. In addition, this information is particularly relevant since it describes the type of islets that are being used in clinical transplantation.

Islets of Higher Mammals from Different Regions of the Pancreas have Different Islet Cell Composition Malaisse-Lagae and colleagues noticed rather signiﬁcant variation in beta and alpha cell populations between PP-rich and PP-poor regions of the human pancreas [24]. Another study by Gersell and colleagues found that a similar difference in islet architecture exists in the canine pancreas [25]. Heterogeneity of endocrine cell types in canine islets was also described by Redecker and colleagues using semithin sections (up to 1000 sections/islet) to reconstruct the islets [16]. A similar approach was used by Baetens and colleagues who examined a large number of serial histological sections (up to 77 sections/ islet), reconstructed individual rat islets, and examined their endocrine cell populations [23]. In contrast to the ﬁndings in the canine and human pancreas and with isolated human islets, the population of beta cells in the rat islets located either in ventral (PP-rich) or dorsal (PP-poor) pancreas was very similar (82.5% versus 82.0%). There was some variation of the alpha cell population in the rat pancreas. The mouse islets described in this chapter appear quite similar to this description of rat islets. Orci and colleagues have suggested that endocrine cell populations in the human pancreas may change throughout life [24,26] with more delta cells and fewer PP cells in the PP-rich region and fewer beta cells in the PP-poor region in the infant pancreas. This

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has not been studied in other species. Thus, in the human pancreas, islets in the uncinate process or the posterior head of the pancreas are greatly enriched in PP-expressing cells, but are relatively “poor” in insulin- and glucagonexpressing islet cells. In contrast, the body and the tail of the pancreas have predominantly insulin- and glucagon-expressing islet cells and relatively few PP-expressing cells.

Islet Interspecies Differences Appear to Impact Islet Function Very little is known about the consequences of the unique endocrine cell organization and cellular composition of islets from higher mammals and whether this affects islet function. Recently, Cabrera and colleagues measured intracellular calcium concentrations, [Ca2+]i, in isolated human islets and found that beta cell oscillatory activity was not coordinated throughout the human islets as it was in mouse islets [14]. Their work also indicated that human and non-human primate islets have a lower glucose threshold for an increase in [Ca2+]i response compared to mouse islets.

Conclusions Overall, the combined data from conventional immunohistochemistry and confocal microscopy indicates that islet architecture and cell composition in higher mammalian species (canine, non-human primate, human) are quite different in the following ways: (1) human islets lack the core–mantle appearance seen in rodent islets; (2) human islets have fewer beta cells and more alpha cells than rodent islets and there is more inter-islet variation in these cell populations in human islets; (3) human along with canine and porcine pancreata have regions where there are islets with considerably more PPexpressing cells. There is little data about differences in islet function between human and rodent islets, but a recent report suggests that these differences in islet architecture and cell composition are associated with differences in islet cell function. The differences in islet architecture and composition between rodents and higher mammals suggest that the transcription factors and extracellular signals controlling islet development and structure may be different in rodents and humans. Acknowledgments. Work in the authors’ laboratory is supported by a Merit Review Award from the VA Research Service, research grants from the National Institutes of Health (DK69603, DK68764, DK66636, DK 68854, DK63439, DK62641), and the Juvenile Diabetes Research Foundation

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M. Brissova and A.C. Powers

International, the Vanderbilt Mouse Metabolic Phenotyping Center (DK59637), and the Vanderbilt Diabetes Research and Training Center (NIH DK20593).

References 1. Bonner-Weir S (1991) Anatomy of islet of Langerhans. In: Samols E (ed) The endocrine pancreas. Raven, New York, pp 15–27 2. Lifson N, Lassa CV, Dixit PK (1985) Relation between blood ﬂow and morphology in islet organ of rat pancreas. Am J Physiol 249:E43–48 3. Bonner-Weir S (1993) The microvasculature of the pancreas, with emphasis on that of the islets of Langerhans. In: Go VLW, DiMagno ER, Gardner JD, Lebenthal FP, Reber HA, Scheele GA (eds) The Pancreas: Biology, Pathobiology, and Disease, 2nd edn. Raven, New York, pp 759–768 4. Prado CL, Pugh-Bernard AE, Elghazi L, Sosa-Pineda B, Sussel L (2004) Ghrelin cells replace insulin-producing beta cells in two mouse models of pancreas development. Proc Natl Acad Sci USA 101:2924–2929 5. Heller RS, Jenny M, Collombat P, Mansouri A, Tomasetto C, Madsen OD, Mellitzer G, Gradwohl G, Serup P (2005) Genetic determinants of pancreatic epsilon-cell development. Dev Biol 286:217–224 6. Wierup N, Sundler F (2005) Ultrastructure of islet ghrelin cells in the human fetus. Cell Tissue Res 319:423–428 7. Wierup N, Yang S, McEvilly RJ, Mulder H, Sundler F (2004) Ghrelin is expressed in a novel endocrine cell type in developing rat islets and inhibits insulin secretion from INS-1 (832/13) cells. J Histochem Cytochem 52:301–310 8. Wierup N, Svensson H, Mulder H, Sundler F (2002) The ghrelin cell: a novel developmentally regulated islet cell in the human pancreas. Regul Pept 107:63–69 9. Bauer GE (1988) Islets of Langerhans. In: Weiss L (ed) Cell and tissue biology, a textbook of histology. Urban & Schwarzenberg, Baltimore, pp 738–749 10. Masharani U, Karam JH, German MS (2001) Pancreatic hormones and diabetes mellitus. In: Greenspan FS, Gardner DG (eds) Basic and clinical endocrinology. Lange Medical Books/McGraw-Hill, New York, pp 658–746 11. Orci L, Baetens D, Rufener C, Amherdt M, Ravazzola M, Studer P, Malaisse-Lagae F, Unger RH (1976) Hypertrophy and hyperplasia of somatostatin-containing D-cells in diabetes. Proc Natl Acad Sci USA 73:1338–1342 12. Dubois M, Pattou F, Kerr-Conte J, Gmyr V, Vandewalle B, Desreumaux P, Auwerx J, Schoonjans K, Lefebvre J (2000) Expression of peroxisome proliferator-activated receptor gamma (PPARgamma) in normal human pancreatic islet cells. Diabetologia 43:1165–1169 13. Brissova M, Fowler MJ, Nicholson WE, Chu A, Hirshberg B, Harlan DM, Powers AC (2005) Assessment of human pancreatic islet architecture and composition by laser scanning confocal microscopy. J Histochem Cytochem 53:1087–1097 14. Cabrera O, Berman DM, Kenyon NS, Ricordi C, Berggren PO, Caicedo A (2006) The unique cytoarchitecture of human pancreatic islets has implications for islet cell function. Proc Natl Acad Sci USA 103:2334–2339 15. Brelje TC, Scharp DW, Sorenson RL (1989) Three-dimensional imaging of intact isolated islets of Langerhans with confocal microscopy. Diabetes 38:808–814 16. Redecker P, Seipelt A, Jorns A, Bargsten G, Grube D (1992) The microanatomy of canine islets of Langerhans: implications for intra-islet regulation. Anat Embryol (Berl) 185:131–141

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17. Stefan Y, Orci L, Malaisse-Lagae F, Perrelet A, Patel Y, Unger RH (1982) Quantitation of endocrine cell content in the pancreas of nondiabetic and diabetic humans. Diabetes 31:694–700 18. Rahier J, Goebbels RM, Henquin JC (1983) Cellular composition of the human diabetic pancreas. Diabetologia 24:366–371 19. Clark A, Wells CA, Buley ID, Cruickshank JK, Vanhegan RI, Matthews DR, Cooper GJ, Holman RR, Turner RC (1988) Islet amyloid, increased A-cells, reduced B-cells and exocrine ﬁbrosis: quantitative changes in the pancreas in type 2 diabetes. Diabetes Res 9:151–159 20. Shapiro AM, Lakey JR, Ryan EA, Korbutt GS, Toth E, Warnock GL, Kneteman NM, Rajotte RV (2000) Islet transplantation in seven patients with type 1 diabetes mellitus using a glucocorticoid-free immunosuppressive regimen. N Engl J Med 343:230–238 21. Street CN, Lakey JR, Shapiro AM, Imes S, Rajotte RV, Ryan EA, Lyon JG, Kin T, Avila J, Tsujimura T, Korbutt GS (2004) Islet graft assessment in the Edmonton Protocol: implications for predicting long-term clinical outcome. Diabetes 53:3107–3114 22. Sakuraba H, Mizukami H, Yagihashi N, Wada R, Hanyu C, Yagihashi S (2002) Reduced beta-cell mass and expression of oxidative stress-related DNA damage in the islet of Japanese Type II diabetic patients. Diabetologia 45:85–96 23. Baetens D, Malaisse-Lagae F, Perrelet A, Orci L (1979) Endocrine pancreas: threedimensional reconstruction shows two types of islets of Langerhans. Science 206: 1323–1325 24. Malaisse-Lagae F, Stefan Y, Cox J, Perrelet A, Orci L (1979) Identiﬁcation of a lobe in the adult human pancreas rich in pancreatic polypeptide. Diabetologia 17:361–365 25. Gersell DJ, Gingerich RL, Greider MH (1979) Regional distribution and concentration of pancreatic polypeptide in the human and canine pancreas. Diabetes 28:11–15 26. Orci L, Stefan Y, Malaisse-Lagae F, Perrelet A (1979) Instability of pancreatic endocrine cell populations throughout life. Lancet 1:615–616 27. Brissova M, Fowler MJ, Wiebe P, Shostak A, Shiota M, Radhika A, Lin PC, Gannon M, Powers AC (2004) Intra-islet endothelial cells contribute to revascularization of transplanted pancreatic islets. Diabetes 53:1318–1325

2. Transcriptional Regulation of Insulin Gene Expression Isabella Artner and Roland Stein

Summary. The insulin gene is expressed exclusively in the beta cells of the islet of Langerhans. The release of this polypeptide hormone into the bloodstream, principally in response to elevated glucose levels, is essential for controlling carbohydrate metabolism in peripheral tissues. A fundamental cause of diabetes, a disease that affects millions of people and is a major cause of morbidity and mortality, is the inability of beta cells to produce sufﬁcient amounts of insulin, resulting in hyperglycemia. A large effort is underway to identify and characterize the transcriptional regulators of genes, like insulin, that are important in islet beta cell function. It is hoped that this knowledge will provide information into how beta cell function is disrupted in type 2 diabetic individuals, and to provide a foundation for cell-based therapies that may be effective in diabetes treatment. Many of the cis-acting sequences, essential in directing both selective and glucose-inducible transcription within the 5′-ﬂanking region of the insulin gene, have been deﬁned and several of the key trans-activators isolated, including PAX-6, PDX-1, MafA, and BETA2/NeuroD1. In addition, the inactivation of genes encoding these regulatory proteins in mice has established that most play a role in islet cell differentiation during pancreas development. In this review, the regulatory role of the islet-enriched transcription factors of the insulin gene will be discussed, with a focus on their role in adult beta cell function.

Introduction Insulin is a powerful regulator of metabolism. This hormone, which is produced by the beta cells of the endocrine pancreas, increases the storage of glucose, fatty acids, and amino acids by its actions on liver, adipose tissue, Department of Molecular Physiology and Biophysics, Vanderbilt University Medical Center, 723 Light Hall, Nashville, TN 37232, USA

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and muscle. The endocrine pancreas is comprised of discrete cellular islands (termed the islets of Langerhans), which are dispersed throughout the larger mass of the exocrine pancreas. The islets of rodents are mainly composed of beta cells, but at their periphery contain four other endocrine cell types, alpha, delta, epsilon, and pancreatic polypeptide (PP) that synthesize the hormones glucagon, somatostatin, ghrelin, and PP, respectively. The primary regulator of insulin expression in pancreatic beta cells is the concentration of blood glucose. The phosphorylation of glucose by glucokinase, which exhibits a high Km for glucose, appears to function as the glucose sensor by adjusting the metabolic ﬂux through glycolysis to the extracellular glucose concentration [1]. Glucose metabolism in beta cells generates intracellular signals which stimulate insulin secretion, insulin mRNA translation, and insulin gene transcription (reviewed in [2]). Diabetes mellitus, the primary human disease affecting the endocrine pancreas, results either from the autoimmune destruction of beta cells (type 1 diabetes) or from defects in the production of insulin and/or sensing of this hormone by peripheral tissues (type 2 diabetes). Because of reduced insulin action, blood glucose levels remain elevated and cause early morbidity and mortality. Despite insulin’s availability and advances in bioengineering that have led to improved technologies for insulin administration, the available systems incompletely mimic the beta cell’s ability to sense and reduce circulating glucose levels sufﬁciently to prevent the severe complications associated with the disease, which predominately targets the kidney, vasculature, and eye. Type 2 is the most common form of diabetes mellitus and largely afﬂicts middle-aged individuals. Genetic studies suggest a major genetic component, but few susceptibility genes have been identiﬁed [3], except in maturity onset diabetes of the young (MODY). MODY, an autosomal dominant form of type 2 diabetes, is characterized by early-onset (usually <25 years) with a primary defect in insulin secretion [4]. There are six identiﬁed genetic loci associated with MODY. Although the ﬁrst locus discovered was for the gene encoding glucokinase [5], it is striking that all of the other characterized ones represent transcription factors involved in the regulation of essential target genes expressed in pancreatic beta cells, and include Hepatocyte Nuclear Factor 1alpha (HNF), HNF1beta, HNF4alpha, PDX-1, and BETA2/NeuroD1 (reviewed in [6]). Notably, BETA2 [7] and PDX-1 [8] are direct regulators of insulin transcription. These ﬁndings illustrate how the identiﬁcation and characterization of transcriptional regulators of isletenriched products, like insulin, provide insight into the disease process. This chapter focuses on how such islet-enriched transcriptional regulators of the insulin gene impact islet beta cell function (see [2,9] for recent pertinent reviews).

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Insulin Gene Expression The 5′-ﬂanking region of the insulin gene that mediates correct tissue- and cell-type-speciﬁc expression in vivo was deﬁned approximately 20 years ago in transgenic mice [10]. This study was very signiﬁcant in that it established the general location of the transcription factor binding sites that direct betacell-type-speciﬁc expression to between −520 and +1 base pairs (bp) relative to the transcription initiation site. Although this experiment was conducted with rat insulin II gene sequences, the same expression data was obtained with the comparable region of the human insulin gene [11,12]. Together, these results indicated that the sequences conserved between mammalian insulin genes that lie within 350 bp of the transcription start site controlled cell-type-speciﬁc expression. (Most animals have a single copy of the insulin gene, although rodents have two non-allelic insulin genes (I and II) that differ in their number of introns and chromosomal location [13].) Consistent with this hypothesis, human and rat insulin sequences were sufﬁcient in directing beta-cell-selective transgenic expression in mice [14]. To further deﬁne the control properties of these 350 bp of 5′-ﬂanking region sequence, insulin-driven reporter activity has been studied in vitro with primary islet cells, as well as transformed islet beta- and non beta-cell lines. These transfection studies have clearly demonstrated that most of the regulation operates through these conserved sequences. The transcription factors binding within the insulin enhancer, which is located between nucleotides −340 and −91, largely determines the cell-speciﬁc and glucose-regulated expression properties of the insulin gene [15–19]. The activity of the enhancer appears to be regulated by both positive- and negative-acting cellular activities [20]. There are a number of cis- and trans-acting factors associated with betacell-speciﬁc activation of the insulin enhancer (Fig. 1). In addition, insulin secreted from beta cells can promote insulin transcription in an autocrine manner by inﬂuencing enhancer-mediated activation [21,22]. Insulinregulated transcription appears to be controlled by the same constellation of factors involved in glucose-regulated expression [21,22], and is mediated in vitro by the insulin receptor [23]. Support for an autoregulatory role for insulin in vivo was also provided upon showing marked abnormalities in glucose homeostasis and beta cell function in mice that have either generally lost insulin receptor substrate-2 expression [24] or speciﬁcally lack the insulin and insulin growth factor-1 receptors [25]. These mice exhibit marked abnormalities in glucose homeostasis and beta-cell function. The cis- and trans-acting factors that are discussed in this review are signiﬁcant effectors of insulin transcription, as concluded from experiments performed in transfected cells. However, insulin transcription is detected in

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USF

PDX-1

PDX-1

+1

PDX-1 MafA

-350

Human Ins C2/A5 E4

Pal E3a E3b

E2

A3

GG2 GG1 C1 (A2)

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Nkx2.2 MafA

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Rat Ins I C2

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A1

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+1

PDX-1

Nkx2.2 MafA

-350

Rat Ins II C2

E2

A3

GG2

C1

E1

A1

TATA

Fig. 1. Schematic of 5′ enhancer region of the human, rat I, and rat II insulin genes. The difference in transcription factor occupancy within the enhancer (i.e., ∼−340/−91) reﬂects sequence divergence between species. The common binding factors are discussed herein, while HNF1alpha [113] and Nkx2.2 [39,66] represent rodent speciﬁc factors

mice lacking key islet-enriched activators (i.e. PAX6 [26], PDX-1 [27], BETA2/ NeuroD1 [28], and MafA [29]). Since it is unclear whether basal and/or inducible transcription is compromised in these animals, it is formally possible that these factors are not bona ﬁde insulin regulators. Yet, it is more likely that each are essential, but that low level insulin expression can occur in their absence. Signiﬁcantly, these islet-enriched regulators are required during the formation and/or function of islet beta cells, with many mutated in type II diabetic individuals with MODY [30].

Principal Factors Regulating Insulin Gene Transcription Detailed characterization of the insulin enhancer indicates that activation is predominately mediated by the A, C, and E elements (Fig. 1), whose core binding motifs are found within the transcription unit of all characterized insulin genes [31]. Stimulation by the factors that act upon these sites results from cooperative processes involving factor–factor, factor–coactivator, and factor–basal transcription complex interactions, although the precise mechanisms required are poorly understood.

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C2 Element Knepel et al. [32] ﬁrst showed that C2 binding activity common to alpha, beta, and epsilon cell extracts contributed to insulin, glucagon, and somatostatin transcription. At the time, this shared binding element was termed the pancreatic islet cell enhancer sequence (PISCES). During the characterization of mice homozygous for the Small eye mutant PAX6 allele, it was realized that the C2 element (−317/−311 bp in the rat I insulin gene) was a PAX6 binding site [26]. This indicated that the reduction in islet hormone synthesis in these mice was caused by the PAX6 mutation, which results in the production of a truncated protein lacking the C-terminal transactivation domain. Further analysis demonstrated that the PISCES complex detected with a rat glucagon G3 probe also contained PAX6. In addition, PAX6 speciﬁcally activated rat insulin I- and glucagon-driven reporter expression in a cotransfected nonislet cell line [26]. Together, these results strongly suggested that PAX6 was a common regulator of many islet hormone genes. PAX6 is a member of the Pax transcription factor family, all of which contain a paired box bipartite DNA binding domain. PAX6 also has a homeodomain. Besides PAX6, the pancreas produces the closely related PAX4 paired/homeodomain protein [33]. Both PAX4 and PAX6 can bind to C2 element sequences, although PAX4 represses PAX6-induced transactivation driven by PAX4/6 recognition sequences [34,35]. It is unclear if PAX4 regulates insulin expression in vivo, as PAX4 expression is not detected in adult beta cells and only transiently in insulin producing cells during development (reviewed in [36]).

Z-Element Region The Z control region (−292/−243 bp) appears to be unique to human insulin, functioning both in glucose-inducible transcription in primary islet cells [37] and as a transcriptional repressor in transformed beta cell lines [37,38] and primary ﬁbroblast cells [37]. A glucose-sensitive DNA-binding complex (termed ZaI) was detected with a −287/−271 bp probe (5′ TGCTCTCCTG GAGACAT 3′) in primary islet nuclear extracts [37]. Recent studies indicate that A element (PDX-1)-mediated activation of the human insulin enhancer depends on upstream sequences of the Z region [39] and that activation of the Z region is at least partially mediated through PDX-1 and MafA binding [40].

A Elements Several mutationally sensitive A + T rich elements are found within the conserved control region of the insulin gene. Each of these A elements contain a

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TAAT core, the central DNA binding recognition motif for homeodomain proteins. Transcription factors in this family are well known for their effect in directing embryonic development and determining differentiated cell identity in a wide variety of organisms (for reviews see [41,42]). The homeodomain is a highly conserved 61 amino acid long helix-turn-helix structural unit that mediates DNA binding, and was ﬁrst identiﬁed in homeotic selector genes of Drosophila melanogaster. The high degree of amino acid conservation within speciﬁc portions of many homeodomain proteins allowed the isolation of PDX-1 from islets by degenerate polymerase chain reaction ampliﬁcation [8,43,44], as well as over a dozen cDNAs from other homeobox genes [44,45]. In addition, the homeodomain Cdx2/3 [46] and Isl-1 [47] transcription factors were isolated from insulinoma cDNA libraries by expression cloning with insulin A element probes. Cdx2/3 can bind in vitro to the A3 element and speciﬁcally stimulate insulin-driven reporter activity in transfected non-insulin producing cells [46]. However, Cdx2/3 expression is restricted to alpha cells within the adult pancreas [48], and null mutant mice only have defects in intestine function and not in beta cells [49]. In contrast, the analysis of Isl-1 null mice demonstrated an essential role for this factor in islet formation during development [50], although the inability to detect Isl-1 binding to an A element probe in beta cell extracts raises serious doubts about its importance in insulin transcription [51]. Isl-1 is, however, present in all islet cell types [52], and can activate somatostatin [53], glucagon [54], and islet amyloid polypeptide (IAPP) [55] gene expression in cotransfection assays. PDX-1 is the predominant binding activity detected with insulin A element probes in pancreatic beta cell extracts [8,56,57]. This factor was ﬁrst characterized as an insulin [8,56–58] and somatostatin [43,44] transcription factor. Although the accepted name is PDX-1 in mice and IDX-1 in humans, this factor has also been referred to as STF-1. PDX-1 expression in the adult pancreas is essentially restricted to islet beta cells (∼91%), a small subset of delta cells (∼15%), and at very low levels in exocrine acinar cells [44,58–60]. PDX-1 is involved in the expression of gene products associated with beta cell identity, including insulin, beta-glucokinase [61,62], IAPP [63,64], and glucose transporter type 2 (GLUT2) [65]. PDX-1 binds within the A element region of each gene in chromatin immunoprecipitation assays (ChIP) [66]. In addition, their transcription is induced upon stable expression of PDX-1 in non-beta cell types [61,67]. Activation of the insulin gene in glucose-treated cells is also dependent upon PDX-1 [17], a process involving both regulated translocation from the nuclear periphery to the nucleoplasm [68,69] and recruitment of the chromatin modiﬁcation machinery [70–72]. In fact, PDX-1 likely plays a crucial role in RNA polymerase II elongation in the nucleus through its ability to recruit the methyltransferase Set9, which inﬂuences

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insulin gene structure by modiﬁcation of histone 3 at lysine 4 [73]. Despite the evidence supporting a direct role for PDX-1 in insulin expression, early gestation embryos in PDX-1 null mice contain a transient population of insulin+ cells [27], demonstrating that insulin can be transcribed independently of PDX-1 under certain circumstances. To address the role of PDX-1 in adult islet beta cells, loxP marked PDX-1 was conditionally inactivated in beta cells using an insulin enhancer-driven Cre recombinase transgenic mouse line [74]. Signiﬁcantly, loss of PDX-1 led to decreased numbers of insulin+, GLUT2+, and IAPP+ cells, while the number of glucagon+ cells increased. The PDX-1 deﬁcient mice developed diabetes, presumably due (at least in part) to the combined loss of insulin and GLUT2. These results established PDX-1 as a central regulator of islet beta cell function, through its actions on genes required in sensing and regulating blood glucose levels.

C1/RIPE3b1 Element The C1 element is located within conserved insulin gene sequences located between approximately −118 and −107 bp [75]. Two distinct factor-DNA complexes are formed with the C1 probe, termed RIPE3b1 and RIPE3b2 [75,76]. The RIPE3b1 gel mobility complex is found exclusively in beta cell nuclear extracts [75,76] whereas RIPE3b2-binding activity is present in a variety of cell types [75]. The RIPE3b2 complex is composed of at least three subunits: p58, p62, and p110 [77]. The p110 subunit of RIPE3b2 is Smbp-2, a protein with potential helicase motifs and a transcription activation domain [77]. The RIPE3b2 activator is not believed to contribute to beta-cell-speciﬁc expression of the insulin gene [75,77]. The DNA-binding component of the RIPE3b1 activator was relatively recently isolated and identiﬁed as the basic leucine-zipper transcription factor MafA [78–80]. Several distinct observations linked MafA to insulin C1 activation, including the demonstration that: (1) MafA-recognizing antibodies detected a protein in the C1 activator gel shift complex; (2) MafA activated C1-element driven reporter expression; (3) MafA was expressed in islet beta cells in vivo; and (4) MafA bound within the enhancer region of the endogenous insulin gene in ChIP assays. In addition, ectopic MafA expression in the glucagonoma alphaTC6 cell line and adenoviral overexpression in rat islets augmented endogenous insulin transcription [81,82]. Subsequent studies identiﬁed several additional Maf response elements within the human and rat insulin enhancer, mutation of which reduces insulin-driven reporter activity in beta cell lines [81]. MafA expression is found exclusively in insulin producing cells in developing and adult pancreas, a unique property in relation to all other islet-enriched

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transcriptional activators [81]. MafA also appears to control glucoseregulated and fatty acid-inhibited insulin expression [82–84]. However, MafA deﬁcient animals had no defects in beta cell development, but adult islet cell function was impaired, with partial loss of insulin expression observed [29]. These results suggest a critical role in beta cell maintenance and function in the adult organ, in part due to its actions on insulin transcription [29]. The absence of a developmental phenotype was surprising when considering that MafA is initially expressed during pancreas organogenesis in the wave of insulin+ cells that are destined to populate the islet [81]. However, expression of the closely related MafB in developing insulin+ cells may act in a redundant fashion [85], since this factor is capable of activating insulin C1-mediated transcription [79,82] and binds to the insulin enhancer region during beta cell formation in vivo (unpublished observations). MafB is only found in alpha cells in adult islets where it acts as a positive regulator of glucagon expression [85]. Signiﬁcantly, MafA and MafB appear to be the principal members of the large Maf family inﬂuencing islet function, with very limited c-Maf expression detected in adult islet cells [86].

E Element The insulin enhancer can be separated into two mini-enhancer units, located between nucleotides −247 to −197 (rat insulin I gene [87]) and −126 to −86 (rat insulin II gene [15,75]) which are capable of directing pancreatic betacell-type-speciﬁc expression of transfected reporter genes. The E site (5′GCCATCTG-3′) is the only common control element within the units. (The insulin I gene in rodents has two E elements (−241 to −233 and −112 to −104 bp), while there is only one (at approximately −100 to −91 bp) in other mammals [31].) Functional interactions between the E activator and PDX-1 within the −247 to −197 bp region [58,88] or C1 (presumably MafA) in −126 to −86 bp region [75,89] appear to be essential for transcriptional activation. Site-directed mutagenesis experiments deﬁned the core insulin E element sequence as 5′ CANNTG 3′ [90], a binding motif also found in the heavy-chain immunoglobulin and muscle creatine kinase control elements [91,92]. A common amino acid DNA-binding region was present in the proteins that positively regulate expression of the heavy-chain immunoglobulin and the muscle creatine kinase enhancer elements. It consisted of a helix-loophelix domain (HLH) that is important in protein–protein interactions and a contiguous amino terminal basic region (b) necessary in DNA-protein binding. This motif is common to a number of transcription factors required in cell type determination, including the muscle determination proteins MyoD [93], Myf-5 [94], and myogenin ([95], as well as the proteins of the

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Drosphila achaete-scute complex, which are important in neural development [96]. The E activator is a dimeric complex composed of bHLH factors that are islet enriched (BETA2 [89]/NeuroD1 [97]) and generally distributed (HEB [98] or E12, E47, or E2/5 [75,99,100]). E47, E12, and E2/5 are all encoded by the E2A gene and share large regions of identity, especially within their carboxy terminal bHLH domains [101]. The ability of bHLH proteins to interact with each other was exploited during BETA2/NeuroD1 isolation, with an E47 bHLH region “bait” used to pull out BETA2 from a pancreatic beta cell cDNA library [89]. The composition of the insulin E activator is typical of other tissue-speciﬁc activators of the bHLH class, of which the neuronal [102] and myogenic [103] bHLH activators are the best described. BETA2/NeuroD1 was originally characterized based upon its ability to activate insulin reporter gene transcription in transfected beta cells (i.e. BETA2 [89]), and neurite formation upon ectopic expression in Xenopus embryos (i.e. NeuroD1 [97]). This factor is expressed in the pancreatic islet [28,104], intestine [104], pituitary [105], and a subset of neurons in the central and peripheral nervous system [97]. BETA2 null mice exhibit speciﬁc defects in each of these tissues [106,107], including causing diabetes in mice due to loss of beta cells [28]. However, BETA2deﬁcient mice can survive diabetes and live to adulthood through the process of beta cell neogenesis, depending upon their genetic background [108]. BETA2/NeuroD1 also appears to be important for glucagon [7], secretin [104], and proopiomelanocortin [105] transcription in the islet, intestine, and pituitary gland, respectively. In addition, the E activator element mediates glucose-inducible transcription of the insulin gene [18,109].

Synergistic Activation Islet-enriched activator binding promotes the assembly of the insulin transcription complex. This process is mediated by interactions between the activators themselves and by contact with the RNA polymerase II apparatus and the chromatin remodeling machinery, or indirectly through bridging coactivators. Thus, PDX-1 and BETA2 binding to the p300 coactivator or its paralogue p300/cAMP response element binding protein-binding protein (CBP) provides a docking and recruitment interface with the general transcriptional machinery [73,110–112]. Individual expression of PDX-1 and BETA2 lead to little or no activation of insulin enhancer-driven expression in non-beta cell lines transfection assays, while MafA alone did so modestly [82]. Combining MafA with PDX-1 or BETA2 produced synergistic activation, with higher activity found when all three proteins were present. MafA was also found to interact with

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PDX-1

MafA

BETA2/ E47

PDX-1 +1 Insulin

p300/CBP

Fig. 2. Islet-enriched activator binding promotes the assembly of the insulin transcription complex. This process is mediated by interactions between activators themselves and by contact with the RNA polymerase II transcriptional apparatus, or indirectly through bridging coactivators. Thus, PDX-1 and BETA2 binding to the p300/CBP coactivators provides a docking and recruitment interface with the general transcriptional machinery. MafA functionally interacts with PDX-1 and BETA2 to promote synergistic activation of the insulin enhancer, although p300/CBP is not involved in this response

endogenous PDX-1 and BETA2 in co-immunoprecipitation assays, while both dominant-negative variants of MafA and small interfering RNAs drastically reduced insulin-driven expression in beta cells. However, p300/CBP was not involved in MafA activation in beta cell lines. These results suggested that interactions between the principal insulin activators MafA, PDX-1, and BETA2 are important for high level expression of the insulin gene in islet beta cells (Fig. 2), a result consistent with the phenotype of MafA null mice [29].

Future Perspectives Until relatively recently, very little was known about the transcription factors that control pancreas development and islet beta cell function. However, with the identiﬁcation and molecular characterization of the insulin transcription factors PDX-1, PAX6, BETA2/NeuroD1, and MafA, our understanding of the mechanisms that control gene expression within the pancreas has increased greatly. The gene knockouts performed on these factors and other pancreasenriched factors has provided insight into key events occurring during pancreatic morphogenesis. Clearly, it would be simplistic to assume that these are the only transcription factors contributing to this process. A continued effort to identify and analyze the function of candidate genes by knockout analysis will be very informative, especially using tissue-speciﬁc

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and inducible knockout strategies to more precisely evaluate the importance of potential regulators in islet cell formation and function. Another important objective will be to determine whether the genes that regulate islet development and insulin biosynthesis contribute to the pathogenesis of diabetes. Because ﬁve of the six characterized MODY genes are transcription factors (PDX-1, HNF1alpha, HNF1beta, HNF4alpha, and BETA2), it is likely that other islet-enriched transcription factors may contribute not only to MODY but to the greater type 2 diabetes population. Finally, if beta cell replacement is to be utilized in the treatment of type 1 and type 2 diabetes, then a comprehensive understanding of islet-enriched transcription factor actions in human beta cell differentiation and function will be essential for developing therapeutic materials. Acknowledgments. Work in the laboratory is supported by National Institutes of Health (RO1 DK49852, DK-50203, DK42502, DK-03-021), American Diabetes Association (7-04-RA116), and Juvenile Diabetes Research Foundation (Advanced Postdoctoral 10-2006-5 to I.A.).

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3. The Biosynthesis of Insulin Donald F. Steiner

Summary. Insulin is synthesized in the pancreatic beta cell via a series of precursor proteins which include preproinsulin and proinsulin. Preproinsulin carries additional information to target the nascent protein chain into the endoplasmic reticulum (ER) where, after cleavage of the signal peptide, it folds efﬁciently to assume the native proinsulin structure stabilized by three disulﬁde bonds. The beta-cell ER appears to be especially adapted to supporting these processes in the face of varying demands for the hormone. However, sustained high-level stimulation of insulin biosynthesis, as in diabetes, may result in beta-cell damage or death via ER stress mechanisms. Correctly folded proinsulin is transferred to the Golgi apparatus from which it is efﬁciently sorted into secretory vesicles of the regulated pathway, where it is converted to insulin and C-peptide. These peptides, along with others such as Islet Amyloid Polypeptide (IAPP/amylin) and various other granin family peptides, are all stored in highly organized mature secretory vesicles/granules, awaiting their regulated discharge into the bloodstream on demand. More information is needed on many aspects of insulin biosynthesis, particularly its regulatory mechanisms and the pathological processes that inﬂuence their function.

Introduction Insulin (Fig. 1) was the ﬁrst protein whose primary structure was elucidated. This feat was accomplished in the mid 1950s by Fred Sanger and led to a Nobel Prize [1]. The structure immediately prompted speculation on the mechanism of assembly of the two-chain hormone, which was not resolved until the discovery of the single-chain precursor molecule in 1967 [2]. This Department of Biochemistry and Molecular Biology, The University of Chicago, 5841 S. Maryland Avenue, MC 1027, Rm N216, Chicago, IL 60637, USA

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Fig. 1. Covalent structure of human insulin. Boxed residues are highly conserved in all known vertebrate insulins. Reproduced from Steiner et al. [97] with permission

discovery and the work that followed revealed a new paradigm for the biosynthesis of biologically active peptides, which eventually led to the broad present-day ﬁeld of protein precursor processing. Post-translational cleavage is now recognized as a widely occurring mechanism for coordinating the biosynthesis and function of many classes of cellular and/or viral proteins. The discovery of proinsulin also revitalized the ﬁeld of insulin research, leading to new approaches to making purer and less immunogenic therapeutic insulins, but also opening new vistas for research in beta-cell biology that continue to expand in today’s powerful genome-oriented research environment.

Structure and Functions of Insulin Precursors Insulin is synthesized via a series of single chain molecules, including preproinsulin, proinsulin and several intermediate cleavage products [3,4] (Fig. 2). The initial translation product of insulin mRNA is preproinsulin, which contains a hydrophobic N-terminal 24 residue signal peptide, a characteristic feature of almost all secreted proteins of animal, plant or bacterial origin. The signal peptide interacts with the signal recognition particle (SRP) [5], a ribonucleoprotein particle in the cytosol, which facilitates the segregation of the nascent proinsulin polypeptide chain from the cytosolic compartment, where protein synthesis is initiated, into the secretory pathway [6,7]. The nascent preprohormone is then translocated via a peptide-conducting channel across the membrane of the rough endoplasmic reticulum (RER) into its lumen. The signal sequence is cleaved by a specialized signal peptidase located on the inner surface of the RER membrane and then rapidly degraded [8]. Within the RER, proinsulin folds and undergoes rapid formation of its three disulﬁde bonds to achieve its native structure [9]. This process is catalyzed by various

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Fig. 2. Covalent structure of human proinsulin. Residues enclosed in dashed circles are the basic amino acid pairs removed during the conversion of proinsulin to insulin (above) and C-peptide (below). Reproduced from Oyer et al. [99] with permission

chaperone proteins, including the enzyme protein-thiol reductase, a resident protein of the endoplasmic reticulum (ER) having a C-terminal KDEL (Lys-Glu-Asp-Leu) ER-retention sequence [10]. Exit from the ER is usually a rate-limiting step in the intracellular transport of many secretory proteins [11], but proinsulin is one of the more rapid. Within 10–20 min it is transported to the Golgi apparatus for further processing and packaging, as described below. After proinsulin is transported through the Golgi apparatus and into immature secretory vesicles, it is cleaved to yield insulin and the 31-residue C-peptide which lacks the basic residue pairs that link it at either end to the B and A chains. Insulin and the C-peptide are stored together in the secretion granules along with small amounts of intact proinsulin and its intermediate cleavage forms [12] as well as islet amyloid polypeptide (IAPP or amylin) and other less abundant beta-cell secretory products [13]. In this respect, proproteins differ from the zymogen forms of many enzymes which are activated after secretion. Intracellular proprotein processing is thus an important feature of almost all peptide-producing endocrine or neural cells, but also occurs in many other tissues as well [3]. Proinsulins vary somewhat in length among species due to variation in the lengths of the connecting polypeptide (C-peptide) which links the C-terminus of the insulin B chain to the N-terminus of the insulin A chain [14]. Most mammalian proinsulins have a pair of basic residues at either end of the Cpeptide, Arg–Arg at the B-chain junction and Lys–Arg at the A-chain junction

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[15,16]. These residues are excised during the conversion of proinsulin to insulin [17,18], resulting in native insulin and the C-peptide. Proinsulin-like molecules in mollusks and insects also have C-peptides roughly similar in size to those in vertebrates and are processed similarly [19,20]. Proinsulin is remarkably similar to insulin in many of its properties, including solubility, isoelectric point [12], self-associative properties [21], and its reactivity with insulin antisera [2,22]. Evidence from nuclear magnetic resonance (NMR) and other structural studies indicate that the conformation of the insulin moiety in proinsulin is nearly identical to that of insulin itself [12,23]. The connecting peptide does not prevent proinsulin from selfassociating to form dimers and zinc-stabilized hexamers [24,25]. The typical length of the C-peptide (30–35 amino acids (aa)) is necessary for efﬁcient processing of proinsulin by the prohormone convertases [26] and also may enhance its folding and intracellular transport. The long redundant connecting segment only partially masks the receptor binding region, i.e., intact proinsulin exhibits 3%–5% of the receptor binding and biologic activity of insulin in vitro [12,27]. Secreted proinsulin and/or intermediate forms do not undergo further cleavage in the circulation or in tissues [28]. Although some crystalline forms of proinsulin have been generated, these have not yielded a three-dimensional structure for the C-peptide domain of the prohormone [29,30]. Thus this domain does not have an ordered structure, as conﬁrmed more recently by NMR studies of proinsulin in solution [23]. Conversion of proinsulin to insulin usually begins about 20 min after synthesis of the preproinsulin chain and continues for 1–2 h [4,31,32] independent of further protein synthesis [33]. As intracellular processing proceeds, intermediate cleavage products such as des-31,32 proinsulin or des-64,65 proinsulin are generated. In man des-31,32 proinsulin is the predominant intermediate form [28,34,35]. The major storage repository is the rich population of mature secretory granules distributed throughout the beta cell cytoplasm. These contain only small amounts (1%–2%) of proinsulin and such intermediate materials and they turn over very slowly over periods of hours to days. Consequently, secreted insulin normally contains only small amounts of these precursor-related peptides. Newly synthesized insulin is selectively released to a slight extent, but the bulk of the secreted material consists of stored preformed hormone and C-peptide [32,36].

Cell Biology of Insulin Biosynthesis The beta cells of the islets of Langerhans share many features with other neurosecretory cells (Fig. 3). The Golgi apparatus plays a critical role in the formation of dense-core secretory granules in these cells [37]. Various post-

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Fig. 3. Summary of the ﬂow (arrows) of (pro)insulin through the secretory pathway of the pancreatic beta cell. Downward dashed arrow indicates mannose-6-P receptormediated sorting pathway of lysosomal hydrolases. Upward dashed arrow indicates autophagic pathway of granule turnover. See text for further details. Reproduced from Steiner et al. [97] with permission

translational modiﬁcations occur in the medial and trans Golgi compartments, such as glycosylation, sulfation, and in some instances, phosphorylation [38]; proinsulin, however, is not known to be modiﬁed in this compartment. The dynamic nature of the Golgi apparatus and the mechanisms that underlie the transport and sorting of secretory products within it are still only incompletely understood [39]. Recent studies favor a model for Golgi progression in which individual cisternae migrate from cis to trans with their secretory contents while small, coated vesicles retrotransport various Golgi resident proteins/enzymes from trans to cis [40]. In this view, secretory products are retained within the same compartment as they mature and ultimately dissipate in the trans Golgi network (TGN) into immature secretory granules. Many of the small Golgi-associated vesicles are coated with a protein called COPI, which is believed to play a major role in directing protein trafﬁc through the Golgi in both anterograde and retrograde directions [41]. Immunocytochemical studies have demonstrated that newly formed clathrin-clad granules

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in the trans Golgi cisternal network (TGN) are rich in proinsulin, conﬁrming that conversion to insulin occurs principally during the maturation of these secretory “progranules” [42]. Energy poisons or Brefeldin A block the transfer of newly formed proinsulin into secretory vesicles, thus preventing its proteolytic processing [42–45]. However, once newly synthesized proinsulin has reached the trans Golgi and/or progranules, its conversion to insulin no longer requires energy [44]. This energy requirement appears to be associated with the budding and/or fusion of the small vesicles which transport secretory products from the ER to the Golgi cisternae [46,41]. The conversion of proinsulin to insulin in intact rat islet cells resembles a ﬁrst-order process having a half-time ranging from about 20 min to 1 h in various studies [12]. Peak labeling of proteins in the Golgi apparatus is observed 30–40 min after biosynthetic labeling of islets with tritiated amino acids; relatively little radioactivity remains in this region after 1 h [47]. A similar pattern is observed by means of electron microscopic immunocytochemistry [42]. Thus, although proinsulin conversion may begin in the trans compartment of the Golgi apparatus, it is now well established that it occurs mainly within newly formed secretory vesicles [48,43] as these leave the Golgi region, and that it continues as these vesicles condense and mature biochemically in the cytosol (Fig. 3). Some studies have indicated that maturing secretory granules in beta cells are sites of continued passive sorting of some constituents, especially those excluded from the condensing dense core [49]. Small amounts of newly generated C-peptide derived from proinsulin by proteolytic processing appear to exit immature granules during their maturation, giving rise to a constitutivelike secretion of C-peptide [50]. The t½ for this process is rather slow (approx. 1.5 h) and the amount released is quantitatively small. It would most likely occur under conditions that stimulate insulin biosynthesis and thus be obscured by increased regulated secretion. Other proteins also are partly targeted into this pathway, including procathepsins B and L [51]. Some furin in the TGN also enters immature secretory granules and is then retrieved via clathrin-coated vesicles [52], but the extent to which it may participate in prohormone processing is unknown. Thus, the maturing secretory granule may function as partial extension of the TGN allowing further sorting of proteins destined for regulated vs. constitutive and/or lysosomal pathways [53].

Mechanism of Proteolytic Conversion of Proinsulin to Insulin Although trypsin acting alone converts proinsulin to an insulin-like component, the product is desB30 insulin due to tryptic cleavage at lysine B29 to release the tripeptide Ala-Arg-Arg from the initial tryptic conversion product,

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insulin-Arg31-Arg32 [15]. However, when pancreatic carboxypeptidase B is combined with trypsin to rapidly remove C-terminal basic amino acid pairs, this secondary cleavage at lysine B29 is suppressed and the naturally occurring products—C-peptide and native insulin—are generated in essentially quantitative yields [54]. This enzyme combination served as a model for the in vivo process, since it gave rise to the known major products and intermediate forms found in pancreatic extracts [16,17]. It also has been used in the commercial production of recombinant human insulin from proinsulin. Early studies with [3H]arginine-labeled islet secretion granules indicated that these are major sites of proinsulin conversion [18]. Moreover, free arginine was released during conversion rather than basic dipeptides, indicating the participation of both trypsin-like and carboxypeptidase B-like activities during secretory granule maturation. Interestingly the carboxypeptidase-like enzyme CPE, was discovered ﬁrst by Fricker et al. in brein in 1985 [56] and similar activities were detected even earlier in isolated islets [55]. Then in 1988 two calcium-dependent endoproteases with proinsulin converting activity were identiﬁed in rat insulinoma secretory granules by Davidson et al. [57]. Subsequent studies revealed that they were members of a larger family of eukaryotic serine proteases related to subtilisin. The ﬁrst member to be found was the yeast pro-alpha-factor processing protease kex2 (or kexin) and its discovery [58] led to the identiﬁcation and characterization of the much more extensive mammalian family of proprotein or prohormone convertases known as the SPCs or PCs [59] (Fig. 4). It was soon established that two members of this family, PC2 and PC1/3, were both expressed in islet beta cells and corresponded to the Type I and Type II secretory granule proteases described by Davidson et al. [57,59]. PC2 was found to cleave proinsulin mainly at the A-chain junction, while PC1/3 preferentially cleaved at the Bchain junction. Moreover the Type II enzyme, PC2, was found to cleave des31,32 proinsulin much more rapidly than the intact prohormone, suggesting a sequential pathway of cleavage [60] (Fig. 5A). Both PC2 and PC1/3 lack the C-terminal transmembrane domain and cytosolic tail present in several other PC family members, including furin, PC7 and PC5/6B (see Fig. 4). These more ubiquitously expressed convertases are localized in the TGN, cycle to the plasma membrane in constitutive vesicles, and are recycled via endosomal pathways [61]. These members process a great variety of proproteins, including growth factor precursors and/or their receptors (e.g. insulin and IGF1 proreceptors), some matrix metalloproteases, and viral glycoproteins. For further background information, see the reviews in [59,62–64]. Because of their role as convertases in secretory granules, it is not surprising that PC2 and PC1/3 are expressed principally in neuroendocrine tissues, including the islets of Langerhans, pituitary, adrenal medulla, and in many regions of the brain, but not at appreciable levels in most other tissues [62].

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Fig. 4. Schematic representation of the structure of the mammalian prohormone convertase (PC) family of serine proteases. The highly conserved catalytic domain (CAT) is stabilized by the downstream essential P domain (P). The propeptide, which must be removed autocatalytically to activate the enzyme, occupies the strongly electro-negative (–) active site (Pro). After cleavage (scissors) and removal of the prodomain the active site can bind and cleave substrates. The variable downstream region (VAR) is essential for the intracellular routing of the proteases but not required for activity. Reproduced from Zhou et al. [64] with permission

In the islets of Langerhans both convertases are detected by immunostaining in the beta cells [65], while only PC2 is detected in the alpha, delta, and pancreatic peptide (PP) producing cells [66]. In beta cells both PC2 and PC1/3 have been colocalized with proinsulin by immunogold labeling in newly formed secretory vesicles, thus conﬁrming the cosegregation of the convertases and their putative substrates [67,68]. The widespread distribution of PC2 and PC1/3 in neuroendocrine cells and tumors [69], along with their more acidic pH optima, indicates that they are key enzymes in the processing of prohormone and neuroendocrine precursors. The PCs all are synthesized as precursors which are activated by autocatalytic cleavage of their N-terminal prodomains usually as a requirement for exiting the ER [70]. In the TGN the prodomain dissociates to release active convertase and then in some instances, e.g., PC1/3, additional autocatalytic cleavages give rise to C-terminally truncated versions which comprise the major active form in the secretory granules [71]. ProPC2 differs from the

A

B

Fig. 5. A Mechanism of proteolytic conversion of proinsulin to insulin. Ribbon model of proinsulin in a conformation suitable for attachment of convertases to the basic residue pairs (side chains shown) for cleavage. Usually PC1/3 (PC3) cleaves initially after R32 (upper bold arrow) followed by PC2 which then cleaves after R65 (lower bold arrow) to liberate Insulin-R⋅R and C-peptide-K⋅R. The basic residues are rapidly removed by CPE to give the native products, insulin and C-peptide. B Cleavage pathways giving rise to the major proinsulin processing intermediates. With human proinsulin the pathway shown to the right (same as in A) predominates, leading to the accumulation of larger amounts of des-31,32 proinsulin. In the absence of PC1/3 some proinsulin is converted to insulin with the accumulation of greater amounts of des 64,65 proinsulin. See text for further details. Panel A courtesy of Gregory Lipkind; panel B reproduced from Steiner et al. [93] with permission

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others in requiring interaction with 7B2, a small secreted neuroendocrine protein. As a result prodomain cleavage is delayed until proPC2 enters secretory granules resulting in a later appearance of active enzyme [71,72]. Prodomain cleavage can occur in the absence of 7B2 but does not result in active PC2, as conﬁrmed by disruption of the 7B2 gene in mice [73]. Calcium plays an important role in the intracellular transport and processing of proinsulin in the beta cell. Depletion of calcium in the secretory pathway has indicated that it normally enters via the ER and is necessary for the activation, transport and action of PC2 and PC1/3 [74]. In rats and probably also pigs and humans, additional partial cleavages of the C-peptide at internal sites appear to be due to other convertases having speciﬁcity for hydrophobic amino acids [75]. The dog C-peptide is also cleaved internally at a single arginine (residue 8) by PC2 or PC1/3 to produce an N-terminally truncated C-peptide having only 23 residues as the main form [76]. Both PC2 and PC1/3 cleave mainly at K⋅R or R⋅R sites, but the PC family generally prefers substrates having a sequence such as R-X-R-X-K/R-R↓, where upstream residues at the 4th or 6th position either augment or are required for cleavage, e.g., furin. PC2 and PC1/3 activity, e.g. on proinsulin, is also augmented by upstream basic residues at the 4th position [64], but they also occasionally cleave at a single Arg or Lys residue at selected sites. Comparative studies of proinsulin processing in PC2 and PC1/3 null mice have conﬁrmed that while either convertase can act alone to cleave proinsulin at both sites to generate some insulin, their combined action gives rise to the most efﬁcient cleavage, as illustrated in Fig. 5B [77,78]. Studies by Rhodes et al. [60] demonstrated that the des-31,32 proinsulin intermediate is a better substrate for cleavage by PC2 and is itself preferentially generated most efﬁciently by PC1/3. This implies that for most efﬁcient processing PC1/3 action must precede that of PC2 and this also ﬁts with the earlier activation of PC1/3 relative to PC2 in the secretory pathway. These facts explain why disruption of the gene encoding PC1/3 much more drastically impairs proinsulin processing (>85% inhibition) than does inactivation of PC2 (∼30% inhibition) [77,78]. As expected, the des-31,32 proinsulin intermediate tends to be elevated in the PC2 nulls, while the des-64,65 intermediate is increased in the PC1/3 nulls. In man, however, des-31,32 proinsulin is normally quite prominent because human proinsulin has only a single upstream basic residue, at the P4 position in the B-chain–C-peptide junction, which enhances the cleavage rate, but only at this site [77]. On the other hand pro-IAPP, the precursor form of islet amyloid polypeptide, which is coexpressed with insulin in the beta cell, relies to a greater extent on PC2 in its processing than on PC1/3 [79,80]. As mentioned above, removal of the Lys and Arg residues exposed by prior endoproteolytic cleavage by the prohormone convertases is an essential

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function carried out mainly by carboxypeptidase E or H (CPE)—a neuroendocrine exopeptidase having selectivity for C-terminal basic residues [81,56] (Fig. 5). Mice lacking active CPE, the CPEfat mouse, exhibit signiﬁcant defects in processing proinsulin and many other neuroendocrine precursors and in addition to obesity, develop diabetes, especially the males [82]. Additional cellular carboxypeptidases, such as carboxypeptidase D, that also may participate in precursor processing in some tissues have been identiﬁed more recently [83]. Removal of basic residues is not only important for efﬁcient and rapid endoproteolytic processing by the PCs, but also prepares certain neuropeptides for C-terminal amidation, a modiﬁcation that is often essential for their activity and is carried out by PAM, the peptidyl amidating monooxygenase [64].

The Beta-Cell Secretory Granule The segregation of proinsulin into the regulated arm of the secretory pathway is highly efﬁcient, with measured levels of 99% or greater, in the immature granules [84]. However, there is controversy as to how efﬁciently it is retained there and whether processing is necessary for its efﬁcient retention, in view of the demonstrated existence of sorting mechanisms for the reorganization of secretory granule contents at least during their maturation [85]. Mature beta-cell secretory granules contain only small amounts (3%–5%) of unprocessed proinsulin and intermediate products [32], and most of the proteolytic processing normally is completed during the ﬁrst 2–3 h after biosynthesis of the preproinsulin chain [77,78]. However, a large fraction of the mature granules remains in the beta cell and these are available for secretion for many hours up to several days [4,12]. The biochemical mechanisms underlying the acquisition of competence for secretion remain poorly understood. Eventually, excess mature granules are taken up and degraded by autophagy, also known as crinophagy, especially if demand for insulin decreases, as for example during extended fasting [4]. The mature secretory granule contains a central dense core consisting essentially of pure insulin molecules arranged in crystalline-like arrays with spacings closely similar to those in typical rhombohedral insulin crystals. These inclusions are suspended in a less electron-dense aqueous halo that contains various ions and small molecules, as well as the bulk of the soluble free C-peptide in amounts closely similar to the amount of insulin [86]. A variety of other minor secretory proteins and peptides, including various secretogranins, IAPP/amylin, and several others are also present at levels well below 1%. Because of the large numbers of stored insulin granules (estimated at 13 000/cell) the pancreatic islets collectively contain a

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sufﬁcient supply of insulin to control carbohydrate metabolism for several days. Cosecretion of insulin and essentially equivalent molar amounts of C-peptide has provided the basis for an alternative measure of insulin production from the beta cell, the C-peptide radioimmunoassay, as developed and reﬁned by Rubenstein, Polonsky and associates [87,88]. The C-peptide assay is especially valuable when extrinsic insulin is being administered but it also provides a highly reliable index of endogenous insulin production under many other clinical conditions as well (for review see [69]).

The ER: A Central Organ in Beta-Cell Function and Survival In addition to its well-established role in the biosynthesis of a wide variety of secreted and/or membrane-inserted proteins, the ER lumen is also the site of their initial folding and structural organization. This is a complex process that requires the intervention of a variety of auxiliary ER-resident proteins which serve as folding sensors and/or chaperonins, ensuring that proteins achieve their native folded states before exiting the ER [90]. Since many secreted proteins have disulﬁde bonds to stabilize them, additional enzymes must be present to assure a ﬂow of oxidizing equivalents into the ER to convert free cysteine residues into disulﬁde bonds in these proteins [91]. To ensure that misfolded proteins do not exit the ER, a quality control system, the unfolded protein response or UPR, acts to attenuate protein synthesis when unfolded proteins accumulate in the ER to higher than normal levels [90]. A variety of evidence indicates that excessive demands on the protein folding machinery of the ER can lead to ER stress and activate a variety of pathways causing beta-cell malfunction or eventual beta-cell death [92]. Normally glucose and certain other nutrient signals regulate proinsulin biosynthesis via rapid stimulation of translation of insulin mRNA which if prolonged, leads to increased synthesis and stability of insulin mRNA as well [93]. This serves to maintain adequate stores of insulin. If glucose stimulation is prolonged or excessive, however, as occurs in diabetes and states of impaired glucose tolerance, a malfunction of ER and related beta cell stress mechanisms may lead to functional failure of the beta cells. Various mixtures of cytokines also have been shown to lead to altered proinsulin biosynthesis and conversion [94]. A dramatic example of the consequences of ER stress is the induction of diabetes in mice due to a point mutation in one allele of the insulin 2 gene that prevents normal formation of the A7–B7 disulﬁde bond [95]. Replacement of CysA7 with tyrosine prevents normal folding of only about 35% of total proinsulin, but this leads to signs of elevated ER stress, and marked

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reduction of the biosynthesis of normal proinsulin. By the fourth week of life, diabetes develops but at this time the beta-cell population is not decreased. However, at later times, the beta cell mass is also decreased [J. Wang, personal communication]. Further explorations of the mechanisms by which the accumulation of misfolded mutant proinsulin impairs normal beta cell function to cause this autosomal dominant form of diabetes should be of great interest, especially in view of the recent discovery that similar insulin gene mutations are a prominent cause of permanent neonatal diabetes in children [96]. An associated question is whether the biosynthesis and efﬁcient folding of proinsulin in the beta cell requires specialized ER components, i.e., proteindisulﬁde isomerases or other chaperonin-like proteins. Recent studies by Dannies and coworkers have raised this interesting and challenging possibility [97]. In conclusion, much has been learned about the biosynthesis of insulin over the past 40 years since the discovery of proinsulin, but many important questions remain regarding such important aspects as the detailed mechanisms and signal transduction pathways involved in the regulation of preproinsulin biosynthesis, the speciﬁc processes ensuring the correct folding of proinsulin, the features responsible for its transport and efﬁcient sorting into secretory vesicles, and the mechanisms underlying their retention, maturation, and mobilization to release appropriate amounts of insulin to maintain metabolic homeostasis. Acknowledgments. I wish to dedicate this review to the many students, postdoctoral fellows and colleagues who have contributed to this area of research over the years since 1967 (for listing see [98]) and to my early scientiﬁc mentors: Herbert S. Anker (d. 1974), Robert H. Williams (d. 1979), and Earl A. Evans, Jr. (d. 1999), who taught me to think independently and creatively, but carefully. Work from this laboratory has been supported by NIH grants DK 13914 and DK 20595 and by the Howard Hughes Medical Institute.

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6. Chan S, Keim P, Steiner D (1976) Cell-free synthesis of rat preproinsulins: characterization and partial amino acid sequence determination. Proc Natl Acad Sci USA 73: 1964–1968 7. Lomedico PT, Chan SJ, Steiner DF, Saunders GF (1977) Immunological and chemical characterization of bovine preproinsulin. J Biol Chem 252:7971–7978 8. Patzelt C, Labrecque A, Duguid J, Carroll R, Keim P, Heinrikson R, Steiner D (1978) Detection and kinetic behavior of preproinsulin in pancreatic islets. Proc Natl Acad Sci USA 75:1260–1264 9. Huang XF, Arvan P (1995) Intracellular transport of proinsulin in pancreatic beta cells. J Biol Chem 270:20417–20423 10. Munro S, Pelham HRB (1987) A C-terminal signal prevents secretion of luminal ER proteins. Cell 48:899–907 11. Lodish HF (1988) Transport of secretory and membrane glycoproteins from the rough endoplasmic reticulum to the Golgi. J Biol Chem 263:2107–2110 12. Steiner DF, Kemmler W, Clark JL, Oyer PE, Rubenstein A (1972) The biosynthesis of insulin. In: Steiner DF, Freinkel N (eds) Handbook of physiology—Section 7 Endocrinology I. Williams & Wilkins, Baltimore, pp 175–198 13. Nishi M, Sanke T, Nagamatsu S, Bell GI, Steiner DF (1990) Islet amyloid polypeptide. A new beta cell secretory product related to islet amyloid deposits. J Biol Chem 265: 4173–4176 14. Steiner DF (1984) The biosynthesis of insulin: genetic, evolutionary and pathophysiologic aspects. In: Gotshlich EC (ed) The Harvey lectures series 78. Academic Press, New York, pp 191–228 15. Chance RE, Ellis RM, Bromer WW (1968) Porcine proinsulin: characterization and amino acid sequence. Science 161:165–167 16. Nolan C, Margoliash E, Peterson JD, Steiner DF (1971) The structure of bovine proinsulin. J Biol Chem 246:2780–2795 17. Steiner D, Cho S, Oyer P, Terris S, Peterson J, and Rubenstein A (1971) Isolation and characterization of proinsulin C-peptide from bovine pancreas. J Biol Chem 246: 1365–1374 18. Kemmler W, Steiner DF, Borg J (1973) Studies on the conversion of proinsulin to insulin III Studies in vitro with a crude secretion granule fraction isolated from islets of Langerhans. J Biol Chem 248:4544–4551 19. Smit AB, Geraerts WPM, Meester I, van Heerikhuizen H, Joose J (1991) Characterization of a cDNA clone encoding molluscan insulin-related peptide II of lymnaea stagnalis. Eur J Biochem 199:699–703 20. Kawakami A, Iwami M, Nagasawa H, Suzuki A, Ishizaki H (1989) Structure and organization of four clustered genes that encode bombyxin, as insulin-related brain secretory peptide of the silkmoth Bombyx mori. Proc Natl Acad Sci USA 86:6843–6847 21. Frank BH, Veros AJ (1968) Physical studies on proinsulin: association behavior and conformation in solution. Biochem Biophys Res Commun 32:155–160 22. Rubenstein AH, Mako M, Welbourne WP, Melani F, Steiner DF (1970) Comparative immunology of bovine, porcine, and human proinsulin and C-peptides. Diabetes 19: 546–553 23. Weiss MA, Frank BH, Khait I, Pekar A, Heiney R, Shoelson SE, Neuringer LJ (1990) NMR and photo-CIDNP studies of human proinsulin and prohormone processing intermediates with application to endopeptidase recognition. Biochemistry 29:8389– 8401 24. Frank BH, Veros AJ (1970) Interaction of zinc with proinsulin. Biochem Biophys Res Commun 38:284–289

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25. Steiner DF (1973) Cocrystallization of proinsulin and insulin. Nature 243:528– 530 26. Taylor NA, Docherty K (1992) Sequence requirements for processing of proinsulin transfected mouse pituitary AtT20 cells. Biochem J 286:619–622 27. Gliemann J, Sorenson HH (1970) Assay of insulin-like activity by the isolated fat cell method: IV. The biological activity of proinsulin. Diabetologia 6:499–504 28. Given BD, Cohen RM, Shoelson SE, Frank BH, Rubenstein AH, Tager HS (1985) Biochemical and clinical implications of proinsulin conversion intermediates. J Clin Invest 76:1398–1405 29. Blundell T, Wood S (1982) The conformation, ﬂexibility, and dynamics of polypeptide hormones. In: Snell EE, Boyer PD, Meister A, Richardson CC (eds) Annual review of biochemistry, vol. 51. Annual Reviews, Palo Alto, pp 123–154 30. Low BW, Fullerton WW, Rosen LS (1974) Insulin/proinsulin, a new crystalline complex. Nature 248:339–340 31. Steiner DF, Clark JL, Nolan C, Rubenstein AH, Margoliash E, Aten B, Oyer PE (1969) Proinsulin and the biosynthesis of insulin. Rec Prog Horm Res 25:207–292 32. Sando H, Borg J, Steiner DF (1972) Studies on the secretion of newly synthesized proinsulin and insulin from isolated rat islets of Langerhans. J Clin Invest 51: 1476–1485 33. Steiner DF, Cunningham DD, Spigelman L, Aten B (1967) Insulin biosynthesis: evidence for a precursor. Science 157:697–700 34. Sobey WJ, Beer SF, Carrington CA, Clark PMS, Frank BH, Gray IP, Luzio SD, Owens DR, Schneider AE, Siddle K, Temple RC, Hales CN (1989) Sensitive and speciﬁc twosite immunoradiometric assays for human insulin, proinsulin, 65–66 split and 32–33 split proinsulins. Biochem J 260:535–541 35. Sizonenko S, Irminger JC, Buhler L, Deng S, Morel P, Halban PA (1993) Kinetics of proinsulin conversion in human islets. Diabetes 42:933–936 36. Gold G, Gishizky ML Grodsky GM (1982) Evidence that glucose marks cells resulting in preferential release of newly synthesized insulin. Science 218:56–58 37. Farquhar MG, Palade GE (1981) The Golgi apparatus complex (1954–1981) from artifact to center stage. J Cell Biol 91:77s–103s 38. Mains RE, Dickerson IM, May V, Stoffers DA, Perkins SN, Ouaﬁk L, Huster EJ, Eipper BA (1990) Cellular and molecular aspects of peptide hormone biosynthesis. Frontiers in Neuroendocrinology 11:52–89 39. Schekman R, Orci L (1996) Coat proteins and vesicle budding. Science 271: 1526–1533 40. Losev E, Reinke CA, Jellen J, Strongin DE, Bevis BJ, Glick BS (2006) Golgi maturation visualized in living yeast. Nature 441:939–940 41. Orci L, Stamnes M, Ravazzola M, Amherdt M, Perrelet A, Sollner TH, Rothman JE (1997) Bidirectional transport by distinct populations of COPI-coated vesicles. Cell 90:335–349 42. Orci L, Ravazzola M, Amherdt M, Madsen O, Vassalli JD, Perrelet A (1985) Direct identiﬁcation of prohormone conversion site in insulin-secreting cells. Cell 42: 671–681 43. Huang XF, Arvan P (1994) Formation of the insulin-containing secretory granule core occurs within immature β-granules. J Biol Chem 269:20838–20844 44. Steiner DF, Clark JL, Nolan C, Rubenstein AH, Margoliash E, Melani F, Oyer PE (1970) The biosynthesis of insulin and some speculation regarding the pathogenesis of human diabetes. In: Cerasi E, Luft R (eds) The pathogenesis of diabetes mellitus, Nobel Symposium 13. Almqvist and Wiksell, Stockholm, pp 57–80

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45. Howell SL (1972) Role of ATP in the intracellular translocation of proinsulin and insulin in the rat pancreatic beta cell. Nat New Biol 235:85–86 46. Lee MC, Miller EA, Goldberg J, Orci L, Schekman R (2004) Bi-directional protein transport between the ER and Golgi. Annu Rev Cell Dev Biol 20:87–123 47. Orci L, Lambert AE, Kanazawa Y, Amherdt M, Rouiller C, Renold AE (1971) Morphological and biochemical studies of B cells in fetal rat endocrine pancreas in organ culture. Evidence for proinsulin biosynthesis. J Cell Biol 50:565–582 48. Orci L, Halban P, Perrelet A, Amherdt M, Ravazzola M, Anderson RG (1994) pHindependent and dependent cleavage of proinsulin in the same secretory vesicle. J Cell Biol 126:1149–1156 49. Arvan P, Kuliawat R, Prabakaran D, Zavacki AM, Elahi D, Wang S, Pilkey D (1991) Protein discharge from immature secretory granules displays both regulated and constitutive characteristics. J Biol Chem 266:14171–14174 50. Kuliawat R, Arvan P (1992) Protein targeting via the constitutive-like secretory pathway in isolated pancreatic islets: passive sorting in the immature granule compartment. J Cell Biol 118:521–529 51. Kuliawat R, Klumperman J, Ludwig T, Arvan P (1997) Differential sorting of lysosomal enzymes out of the regulated secretory pathway in pancreatic beta cells. J Cell Biol 137:595–608 52. Dittié AS, Thomas L, Thomas G, Tooze SA (1997) Interaction of furin in immature secretory granules from neuroendocrine cells with the AP-1 adaptor complex is modulated by casein kinase II phosphorylation. EMBO J 16:4859–4870 53. Arvan P, Castle J (1992) Protein sorting and secretion granule formation in regulated secretory cells. Trends Cell Biol 2:327–331 54. Kemmler W, Peterson JD, Steiner DF (1971) Studies on the conversion of proinsulin to insulin. I. Conversion in vitro with trypsin and carboxypeptidase B. J Biol Chem 246:6786–6791 55. Zühlke H, Steiner DF, Lernmark A, Lipsey C (1976) Carboxypeptidase B-like and trypsin-like activities in isolated rat pancreatic islets. In: CIBA Foundation (eds) Polypeptide hormones: molecular and cellular aspects. Elsevier Excerpta Medica North-Holland, Amsterdam, pp 183–195 56. Fricker L, Evans C, Esch F, Herbert E (1986) Cloning and sequence analysis of cDNA for bovine carboxypeptidase E. Nature 323:461–464 57. Davidson H, Rhodes C, Hutton J (1988) Intraorganellar calcium and pH control proinsulin cleavage in the pancreatic cell via two distinct site-speciﬁc endopeptidases. Nature 333:93–96 58. Julius D, Brake A, Blair L, Kunisawa R, Thorner J (1984) Isolation of the putative structural gene for the lysine-arginine-cleavage endopeptidase required for processing of yeast prepro-alpha-factor. Cell 37:1075–1089 59. Steiner DF, Smeekens SP, Ohagi S, Chan SJ (1992) The new enzymology of precursor processing endoproteases. J Biol Chem 267:23435–23438 60. Rhodes C, Lincoln B, Shoelson S (1992) Preferential cleavage of des-31,32-proinsulin over intact proinsulin by the insulin secretory granule type II endopeptidase. J Biol Chem 267:22719–22727 61. Molloy S, Thomas L, VanSlyke J, Stenberg P, Thomas G (1994) Intracellular trafﬁcking and activation of the furin proprotein convertase: localization to the TGN and recycling from the cell surface. EMBO J 13:18–33 62. Rouillé Y, Duguay S, Lund K, Furuta M, Gong Q, Lipkind G, Oliva A Jr, Chan S, Steiner D (1995) Proteolytic processing mechanisms in the biosynthesis of neuroendocrine

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76.

77.

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peptides: the subtilisin-like proprotein convertases. Front Neuroendocrinol 16: 322–361 Thomas G (2002) Furin at the cutting edge: from protein trafﬁc to embryogenesis and disease. Nat Rev Mol Cell Biol 3:753–766 Zhou A, Webb G, Zhu X, Steiner DF (1999) Proteolytic processing in the secretory pathway. J Biol Chem 274:20745–20748 Smeekens SP, Montag AG, Thomas G, Albiges-Rizo C, Carroll R, Benig M, Phillips LA, Martin S, Ohagi S, Gardner P, Swift HH, Steiner DF (1992) Proinsulin processing by the subtilisin-related proprotein convertases furin, PC2 and PC3. Proc Natl Acad Sci USA 89:8822–8826 Tanaka S, Kurabuchi S, Mochida H, Kato T, Takahashi S, Watanabe T, Nakayama K (1996) Immunocytochemical localization of prohormone convertases PC1/PC3 and PC2 in rat pancreatic islets. Arch Histol Cytol 59:261–271 Malide D, Seidah NG, Chretien M, Bendayan M (1995) Electron microscopic immunocytochemical evidence for the involvement of the convertases PC1 and PC2 in the processing of proinsulin in pancreatic beta cells. Histochem Cytochem 43:11–19 Itoh Y, Tanaka S, Takekoshi S, Itoh J, Osamura R (1996) Prohormone convertases (PC1/3 and PC2) in rat and human pancreas and islet cell tumors: subcellular immunohistochemical analysis. Pathol Int 46:726–737 Scopsi L, Gullo M, Rilke F, Martin S, Steiner DF (1995) Proprotein convertases (PC1/ PC3 and PC2) in normal and neoplastic human tissues: their use as markers of neuroendocrine differentiation. J Clin Endocrinol Metab 80:294–301 Anderson E, VanSlyke J, Thulin C, Jean F, Thomas G (1997) Activation of the furin endoprotease is a multiple-step process: requirements for acidiﬁcation and internal propeptide cleavage. EMBO J 16:1508–1518 Zhou A, Mains RE (1994) Endoproteolytic processing of proopiomelanocortin and prohormone convertases 1 and 2 in neuroendocrine cells overexpressing prohormone convertases 1 or 2. J Biol Chem 269:17440–17447 Muller L, Zhu P, Juliano MA, Juliano L, Lindberg I (1999) A 36 residue peptide contains all of the information required for 7B2-mediated activation of hormone convertase 2. J Biol Chem 274:21471–21477 Westphal C, Muller L, Zhou A, Zhu X, Bonner-Weir S, Steiner D, Lindberg I, Leder P (1999) The neuroendocrine protein 7B2 is required for peptide hormone processing in vivo and provides a novel mechanism for pituitary Cushing disease. Cell 96: 689–700 Guest PC, Bailyes EM, Hutton J (1997) Endoplasmic reticulum Ca2+ is important for the proteolytic processing and intracellular transport of proinsulin in the pancreatic beta-cell. Biochem J 323:445–450 Verchere C, Paoletta M, Neerman-Arbez M, Rose K, Irminger J, Gingerich R, Kahn S, Halban P (1996) Des-(27–31) C-peptide. A novel secretory product of the rat pancreatic beta cell produced by truncation of proinsulin connecting peptide in secretory granules. J Biol Chem 271:27475–27481 Kwok S, Chan S, Steiner D (1983) Cloning and nucleotide sequence analysis of the dog insulin gene: coded amino acid sequence of canine preproinsulin predicts an additional C-peptide fragment. J Biol Chem 258:2357–2363 Furuta M, Carroll R, Martin S, Swift H, Ravazzola M, Orci L, Steiner DF (1998) Incomplete processing of proinsulin to insulin accompanied by elevation of des-31,32 proinsulin intermediates in islets of mice lacking active PC2. J Biol Chem 273: 3431–3437

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78. Zhu X, Orci L, Carroll R, Norrbom C, Ravazzola M, Steiner DF (2002) Severe block in processing of proinsulin to insulin accompanied by elevation of des-64,65 proinsulin intermediates in islets of mice lacking prohormone convertase 1/3. Proc Natl Acad Sci USA 99:10299–10304 79. Wang J, Xu J, Finnerty J, Furuta M, Steiner DF, Verchere CB (2001) The prohormone convertase enzyme 2 (PC2) is essential for processing of pro-islet amyloid polypeptide at the N-terminal cleavage site. Diabetes 50:534–539 80. Marzban L, Trigo-Gonzalez G, Zhu X, Rhodes CJ, Halban PA, Steiner DF, Verchere CB (2003) Role of beta-cell prohormone convertase 1/3 in processing of pro-islet amyloid polypeptide. Diabetes 53:141–148 81. Guest PC, Rhodes CJ, Hutton JC (1989) Regulation of the biosynthesis of insulinsecretory-granule proteins. Biochem J 257:431–437 82. Naggert J, Fricker L, Varlamov O, Nishina P, Rouille Y, Steiner D, Carroll R, Paigen B, Leiter E (1995) Hyperproinsulinemia in obese fat/fat mice is associated with a point mutation in the carboxypeptidase E gene and reduced carboxypeptidase activity in the pancreatic islets. Nat Genet 10:135–142 83. Song L, Fricker LD (1996) Tissue distribution and characterization of soluble and membrane-bound forms of metallocarboxypeptidase D. J Biol Chem 271:28884– 28889 84. Rhodes CJ, Halban PA (1987) Newly synthesized proinsulin/insulin and stored insulin are released from pancreatic beta cells predominantly via a regulated, rather than a constitutive, pathway. J Cell Biol 105:145:53 85. Arvan P, Halban P (2004) Sorting ourselves out: seeking consensus on trafﬁcking in the beta-cell. Trafﬁc 5:53–61 86. Michael J, Carroll R, Swift H, Steiner DF (1987) Studies on the molecular organization of rat insulin secretory granules. J Biol Chem 262:16531–16535 87. Rubenstein AH, Clark JL, Melani F, Steiner DF (1969) Secretion of proinsulin Cpeptide by pancreatic B cells and its circulation in blood. Nature 224:697–699 88. Van Cauter E, Mestrez F, Sturis J, Polonsky KS (1992) Estimation of insulin secretion rates from C-peptide levels. Comparison of individual and standard kinetic parameters for C-peptide clearance. Diabetes 41:368–377 89. Polonsky KS (1995) The beta-cell in diabetes: from molecular genetics to clinical Research. Diabetes 44:705–17 90. Schröder M, Kaufman RJ (2005) ER stress and the unfolded protein response. Mutat Res 569:29–63 91. Frand AR, Cuozzo JW, Kaiser CA (2000) Pathways for protein disulphide bond formation. Trends Cell Biol 10:203–210 92. Corbett JA (2006) Insulin biosynthesis: The IREny of it all. Cell Metab 4:175–183 93. Steiner DF, Bell GI, Rubenstein AH, Chan SJ (2006) Chemistry and biosynthesis of the islet hormones: insulin, islet amyloid polypeptide (amylin), glucagon, somatostatin, and pancreatic polypeptide. In: DeGroot L, Jameson JL (eds) Endocrinology 5th edn. Saunders, Philadelphia, Chapter 48, pp 925–960 94. Hostens K, Pavlovic D, Zambre Y, Ling Z, Van Schravendijk C, Eizirik DL, Pipeleers DG (1999) Exposure of human islets to cytokines can result in disproportionately elevated proinsulin release. J Clin Invest 104:67–72 95. Wang J, Takeuchi T, Tanaka S, Kubo SK, Kayo T, Lu D, Takata K, Koizumi A, Izumi (1999) A mutation in the insulin 2 gene induces diabetes with severe pancreatic betacell dysfunction in the Mody mouse. J Clin Invest 103:27–37 96. Støy J, Edghill EL, Flanagan SE, Ye H, Paz VP, Piuzhnikov A, Below JE, Hayes MG, Cox NJ, Lipkind GM, Lipton RB, Greeley SA, Patch AM, Ellard S, Steiner DF, Hattersley

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AT, Philipson LH, Bell GI (2007) Insulin Gene Mutations as a Cause of Permanent Neonatal Diabetes. PNAS 104:15040–15044 97. Zhu YL, Abdo A, Gesmonde JF, Zawalich KC, Zawalich W, Dannies PS (2004) Aggregation and lack of secretion of most newly synthesized proinsulin in non- beta-cell lines. Endocrinology 145:3840–3849 98. Steiner DF, Chan SJ, Rubenstein AH (2001) Biosynthesis of insulin. In: Jefferson LS, Cherrington AD (eds) Handbook of physiology—The endocrine system II. Oxford University Press, Chapter 3, pp 49–77 99. Oyer PE, Cho S, Peterson JD, Steiner DF (1971) Studies on human proinsulin: Isolation and amino acid sequence of the human pancreatic C-peptide. J Biol Chem 246:1375– 1386

II. Cell Signaling and Insulin Secretion

4. Metabolic Regulation of Insulin Secretion Barbara E. Corkey

Introduction Understanding fuel-induced insulin-secretion cascades is essential to assessing beta-cell pathology in diabetes. Such understanding may pave the way for more effective therapies to treat or prevent diabetes and to maintain islets or surrogate beta cells for transplantation. Metabolic regulation of insulin secretion requires signals reﬂective of the metabolic state. Signals generally require large rapid changes of limited duration, competitive opposing signals, or ability to translocate. Here we will discuss the metabolic origin and consequences of the following putative metabolic signals that each exhibit such attributes: the adenosine triphosphate/adenosine diphosphate (ATP/ADP) ratio (competitive opposition), malonyl CoA (rapid transient)/ long chain acyl CoA (LC-CoA) (competitive opposition, translocation), the redox state (competitive opposition), and the role of oscillations (rapid transient).

Consensus Model and the Rise of Ca2+ (Fig. 1) The beta cell possesses a unique signal transduction system dependent on metabolism of fuel stimuli to initiate insulin secretion [1,2]. It has been dogma in the ﬁeld that glycolytic and oxidative events leading to accelerated ATP generation are the key transduction phenomena in beta-cell signaling [2,3]. A rise in the cytosolic ATP/ADP ratio is thought to close metabolically sensitive KATP channels depolarizing the beta cell, activating voltage-gated Ca2+ channels and elevating intracellular Ca2+ to promote secretion. Obesity Research Center, Department of Medicine, Boston University School of Medicine, Boston, MA 02118, USA

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Glucose ATP/ADP ↓

Glucose ATP G6P

+

ADP

F6P

O 2 use

+ PFK

-

-

FDP ADP ATP

ATP/ADP ↑

Ca 2 + cyto ↑

Pyr

-

K+

Insulin }

Ca 2+

Fig. 1. Consensus model of glucose induced increase in the adenosine triphosphate/adenosine diphosphate (ATP/ADP) ratio and cytosolic free Ca2+. G6P, glucose 6-phosphate; F6P, fructose 6-phosphate; PFK, phosphofructokinase 1; FDP, fructose 1,6 bisphosphate; Pyr, pyruvate

The ATP/ADP Ratio The power of the cytosolic energy state to regulate metabolism has been known for decades. It is probably the oldest established metabolic signal and is reﬂected in several expressions each of which has a relationship to oxidative phosphorylation: ATP/ADP, free ADP3−, [ATP·AMP]/[ADP]2, or ATP/[ADP·PO4]. For the purposes of this article, we will simplify by using the ATP/ADP ratio. Mitochondrial respiration increases and decreases rapidly in response to decreased and increased energy state of the cytosolic compartment. We have shown that the beta cell responds as expected to changes in the ATP/ADP ratio with increased respiration when the ratio is low and decreased respiration when the ratio is high (Fig. 2). The sensing mechanism to changes in the ATP/ADP ratio is inherent in the adenine nucleotide transporter that senses free adenine nucleotides, ATP4− and ADP3− [4], and regulation of oxidative phosphorylation by ADP and ATP.

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55

O2 Consumption (nmol/min/mg protein)

10

8

6

4

2

0 3

150

15

ATP/ADP

Fig. 2. Relationship between the ATP/ADP ratio and O2 consumption. The ATP/ADP ratio was ﬁxed using creatine, creatine phosphate, creatine phosphokinase, and ATP plus ADP in permeabilized clonal beta cells. O2 consumption was measured in a cell suspension using a Clark type electrode

Table 1. Distribution of adenine nucleotides in isolated hepatocytes and liver mitochondria Total (mM) Free (mM) Mg bound (mM) Cytosol ATP ADP Ratio Mitochondria ATP ADP Ratio

2.76 0.32 8.6

0.16 0.15 1.1

2.60 0.17 15.3

10.4 5.84 1.8

0.44 2.49 0.2

9.88 3.35 2.9

Data are taken from Corkey and colleagues [6]

ATP and ADP exist in cells in multiple forms that are in equilibrium at a given pH and are distributed mainly between cytosolic and mitochondrial compartments. This is illustrated by direct measurements made in liver (Table 1). It should be noted that the free ATP (ATP4−) is a small proportion of the total ATP and the adenine nucleotide translocator is unusual in sensing this form rather than the more common Mg·ATP that is the substrate for ATP-requiring enzymes. It is interesting to note that the ATP-sensitive K+channel (KATP-channel) is the other protein known to require ATP4− [5]. Furthermore, there are large differences in the ratios of different moieties in cytosol and mitochondrial compartments while the whole cell ratio is usually closer to the cytosolic ratio.

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ATP/ADP Measurements in Beta Cells Compartmental distribution of adenine nucleotides has not been done in islets. However, the group of Henquin has performed careful measurements of the ATP/ADP ratio in mouse islets and found a good correlation between the ratio and insulin secretion induced by glucose [7–9]. It has also been demonstrated that the ATP/ADP ratio oscillates in islet cells [10] and clonal beta cells [11]. There has been relatively less focus on the relationship between other fuel secretagogues and the ATP/ADP ratio. Additional work is needed to carefully assess the pattern of ATP/ADP change relative to insulin secretion in response to other stimuli. It is particularly important to do this at early time points and with frequent sampling to assess oscillatory changes [10,12–14].

The ATP/ADP Ratio and O2 Consumption There are several enigmas regarding ATP/ADP control in the beta cell. Stimulation of insulin secretion by fuels is accompanied by a rise in the ATP/ADP ratio that is essential to close the KATP-channel and cause depolarization of the cell. At the same time, it is clear that oxygen consumption increases despite the established ability of a rise in the ATP/ADP ratio to inhibit respiration (Fig. 2). This increase in respiration has been attributed by some to a rise in cytosolic Ca2+. However, Ca2+ does not directly stimulate respiration in beta cells [15], increased respiration precedes the rise in the ATP/ADP ratio, and both precede the rise in Ca2+. Thus, Ca2+ cannot cause increased respiration (Fig. 3). We have proposed that the explanation for this apparent ATP/ADP decrease

<3

NAD(P)H

21±7

O2

29±12

ATP/ADP increase

44±9

FAD +

53±8

Malonyl CoA

< 60

LC-CoA decrease

~120 325±20

Ca2 + 0

100

200

300

Half-maximal change (sec)

400

Fig. 3. Time course of changes following glucose addition to a suspension of clonal beta cells; adenosine triphosphate (ATP), adenosine diphosphate (ADP), malonyl CoA, and long chain acyl-CoA (LC-CoA) were measured enzymatically, O2 was measured using a Clark type electrode, NAD(P)H and FAD+ were measured by their intrinsic ﬂuorescence, and Ca2+ was measured using fura-2

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enigma is that the earliest events in glucose metabolism are phosphorylation by glucokinase and phosphofructokinase using ATP to cause a transient decline in the ATP/ADP ratio to initially stimulate respiration while oscillations in the ratio alternately close the channel and stimulate respiration.

The Lag Between ATP/ADP and Ca2+ Rise A second enigma concerns the reason for the lag in secretion and the events that occur between the rise in the ATP/ADP ratio, and the rise in Ca2+ and the onset of secretion. Whereas the peak in the ATP/ADP ratio occurs by 1 min, the rise in Ca2+ requires at least another 60 s to begin and does not peak until about 4 min. What happens during this period of time? Is it possible that the ATP/ADP ratio does not act alone to close the KATP-channel? The effect of ATP to close the channel is rapid, milliseconds or less, in a cell-detached patch so one cannot infer that the interaction between the channel and ATP is slow. We have hypothesized that the lag is due to the ability of LC-CoA or other lipid metabolites to open the channel, since the decrease in LC-CoA occurs after the rise in the ATP/ADP ratio and appears to occur closer to the time when Ca2+ rises.

Ca2+ Often Does Not Correlate with Secretion The metabolic events discussed up to now concern the steps leading to the rise in Ca2+ that is considered an immediate precursor to exocytosis. The role of Ca2+ and the importance of Ca2+ in insulin secretion has been well documented in the literature [16–21] and will not be discussed here. However, there are many examples of insulin secretion that do not correlate with increases in cytosolic Ca2+ [22–25]. We have recently shown a lack of correlation between insulin secretion and the Ca2+ transient between 8 and 12 mM glucose in mouse islet cells [26]. These studies documented that the maximum Ca2+ response occurred between 4 and 8 mM glucose whereas insulin secretion continued to increase above 8 mM glucose. Under physiological conditions where variable levels of such incretins as acetylcholine, GLP-1, or free fatty acids (FFA) are present, a small increase in glucose causes a much greater increase in insulin secretion than glucose alone, without an increase in Ca2+ levels above that caused by glucose. Henquin and colleagues documented normal concentration dependent glucose stimulated insulin secretion under conditions where Ca2+ levels were elevated but unchanging [27]: the KATP-channel independent pathway. Thus, there must be metabolic signals generated by glucose and perhaps other fuels in addition to Ca2+.

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Lipid Metabolism in the Beta Cell and a Central Role for Cytosolic LC-CoA (Fig. 4) Insulin secretion is essential to store all fuels including FFA, which are the main energy source for the body. Thus, the beta cell must be able to sense the difference between an elevation in glucose and an elevation in both glucose and FFA. Within the beta cell there are also interactions between glucose and FFA, as LC-CoA the activated form of FFA, such that glucose-induced insulin release is associated with inhibition of FFA oxidation and increased lipid synthesis in pancreatic beta cells [28–30]. Indeed, signiﬁcant increases occur in the total mass of diacylglycerol [31] and phosphatidic acid [32,33] in glucose-stimulated beta cells. Glucose and endogenous LC-CoA are the main sources of the glycerol and lipid components, respectively, of diacylglycerol and phosphatidic acid [32,34,35]. There is a well-documented mechanism by which the fate of intracellular FFA is determined, as elegantly demonstrated by McGarry and colleagues [36–41]. His were the ﬁrst demonstrations, in liver, that the fate of FFA was determined by malonyl CoA. We have subsequently provided evidence of

Glucose

Glucose

DHAP PS G3PD Glycerol 3-phosphate

GPAT

ACS FFA

FFA

PI

PC PE

PPH-1 LPA-AT DGAT LPA PA DG TG

LCLC-CoA

Glycerol

FFA

CPT1 Acylcarnitine

b-Oxidation CO2

Fig. 4. Pathway of complex lipid formation. DHAP, dihydroxyacetone phosphate; G3PD, glycerol 3-phosphate dehydrogenase; ACS, acyl CoA synthetase; CPT, carnitine palmitoyl transferase; GPAT, glycerolphosphate acyl transferase; LPA, lysophosphatidic acid; AT, acyl transferase; PA, phosphatidic acid, PI, phosphoinositide; PPH, phosphatidate phosphohydrolase; DG, diacylglycerol, PC, phosphatidylcholine; PE, phosphatidylethanolamine; PS, phosphatidylserine; TG, triglyceride

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similar regulation in beta cells and further hypothesized that the shift from fat oxidation to synthesis generates signals reﬂective of excess fuel and the need for insulin secretion [42–44]. We demonstrated that glucose causes marked alterations in the acyl CoA proﬁle of clonal pancreatic beta cells, with the largest (5-fold) and earliest (by 2 min) change occurring in malonyl CoA [43,44]. Indeed, nutrients or combinations that can both increase the production of acetyl CoA and provide anaplerotic input into the citric acid cycle cause insulin release [45–47]. This is a condition that is essential for the production of the malonyl CoA necessary to elevate cytosolic LC-CoA because efﬂux of the malonyl CoA precursor, citrate, from the mitochondria does not occur unless there has been compensatory input into the citric acid cycle. In addition, exogenous fatty acids potentiate glucose-stimulated secretion [30,48–51], possibly by providing additional acyl groups for LC-CoA or complex lipid synthesis. It should be noted that increases in FFA in the absence of stimulatory glucose do not stimulate secretion, at least in part, because malonyl CoA levels are not elevated. The model derived from these studies (Fig. 5) includes malonyl CoA derived from glucose via glycolysis, anaplerosis via pyruvate carboxylase in the citric acid cycle, and export of citrate from the mitochondria to the cytosol where

FFA

FFA

Complex Lipids

ACS LC-CoA

Glycolysis

Glucose

CPT-1

Malonyl CoA

-

ACC

PD

H

AcCoA

pyruvate

pyruvate PC

AcCoA CS

citrate

CL citrate

OAA

Mitochondria cytosol

Fig. 5. Model of malonyl CoA formation from glucose. ACS, acyl CoA synthetase; CPT, carnitine palmitoyl transferase; PDH, pyruvate dehydrogenase; PC, pyruvate carboxylase; CS, citrate synthase; CL, citrate lyase; ACC, acetyl CoA carboxylase

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it is ultimately converted to malonyl CoA via a pathway regulated by acetyl CoA carboxylase activity. Thus, in the presence of plentiful glucose, malonyl CoA production serves to inhibit FFA oxidation and facilitate glucose oxidation. When FFA oxidation is inhibited by malonyl CoA, cytosolic LC-CoA rises, at least transiently, and conversion to complex lipids occurs (Fig. 4) [43,44]. Free fatty acids, not glucose, are believed to be the major endogenous energy source for unstimulated islets [29,48]. Stimulation of islets by glucose diminishes FFA oxidation and increases total respiration [52]. Thus, one of the metabolic events induced by glucose stimulation appears to be a shift from fatty acids to glucose as an oxidative fuel [15].

LC-CoA as a Signal The only two known effects of malonyl CoA are to act as a precursor of de novo FFA synthesis from glucose, that occurs to a miniscule extent in beta cells [44], and to regulate CPT1. The lack of signaling targets for malonyl CoA suggest that the cytosolic lipid changes that result from its elevation are more likely to play a signaling role. The elevations in LC-CoA, phosphatidic acid, and diacylglycerol resulting from glucose stimulation [31,33] directly activate PKC isoforms [53]. Importantly, LC-CoA inﬂuences the ATP/ADP ratio through modulation of the adenine nucleotide translocase [54–57]. LC-CoA may modulate other enzymes [58,59] and channels [60–64], or modify the acylation state of key proteins involved in regulation of ion channel activity and exocytosis [65,66]. The total CoA pool is ﬁxed, over short intervals, and distributed between mitochondrial and cytosolic compartments that are not interchangeable [67,68]. Thus, the maximum LC-CoA concentration is limited by the total CoA pool in the relevant compartment. High fat and certain drugs or steroids have the potential to increase this pool over a period of hours to days.

LC-CoA as a Signal is Contentious Support for LC-CoA, as the initiator of an acylation cascade, has been discussed by us in numerous reviews [42,43,65,69,70]. In contrast, others have shown that overexpressing malonyl CoA decarboxylase caused a reduction in total cellular malonyl-CoA with unchanged glucose stimulated insulin secretion [71,72]. An important missing piece of information from these studies is assessment of whether early increases in malonyl CoA in response to glucose, albeit from a much lower basal level, were sufﬁcient to stimulate complex lipid synthesis.

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Effect of LC-CoA on Channels The KATP-channel is regulated by several metabolites known to change in response to glucose: ATP4− closes the channel and Mg·ADP and LC-CoA esters both open the channel [5,60–64,73]. It is likely that the balance among these metabolites regulates the metabolic sensitivity of the channel. We have suggested that lipotoxicity may exert some of its negative effects as a consequence of sustained elevations in LC-CoA that could both lower ATP4− levels [56] and diminish the ability of ATP4− to close the channel.

Effect of LC-CoA on Lipolysis It has been reported that LC-CoA inhibits hormone sensitive lipase (HSL) [57]. We have recently conﬁrmed that ﬁnding but surprisingly found that LC-CoA stimulated lipase activity in beta cells even in HSL knockout islets [74], implying that HSL is not the main lipase in beta cells. More importantly, the ability of glucose to stimulate lipolysis [75] may result from glucoseinduced inhibition of FFA oxidation by malonyl CoA with a resulting increase in cytosolic LC-CoA. Stimulation of lipolysis by glucose provides an additional mechanism to increase LC-CoA.

Effect of LC-CoA on Exocytosis It has been reported that LC-CoA is essential for exocytosis in some cells [76]. Deeney and colleagues showed, in permeabilized clonal beta cells, that LC-CoA directly stimulated exocytosis only when Ca2+ was elevated [77]. They also showed that an increase in membrane capacitance, indicative of granule fusion with the plasma membrane, occurred in response to LCCoA and that this could be blocked by cerulenin, an acylation inhibitor [77].

Mitochondrial Signals, Redox, and Energy Metabolism in the Beta Cell In most mammalian cells, 90% of the overall metabolic rate is due to mitochondrial O2 consumption, most of which is coupled to ATP production. Mitochondria play an important role in fuel-induced insulin secretion [13,42,78–84], although the precise nature of that role is not clear. It is well established that interfering with mitochondrial metabolism inhibits insulin

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B.E. Corkey

Mitochondria Succ

Cytosol Succ

Dicarboxylate carrier

Mal

Mal Pyr

Pyr

Pyruvate carrier

OH Mal

OH NAD

NADP

MDH

OAA

Mal ME

NADH NADPH

Pyr

PC Pyr Fig. 6. Carriers and reactions involved in succinate entry into the mitochondria. Succ, succinate; Mal, malate; Pyr, pyruvate; MDH, malate dehydrogenase; ME, malic enzyme; OAA, oxaloacetate

secretion from glucose and other fuels [15,85]. In addition, there are fuels that generate secretory signals exclusively through mitochondrial metabolism such as methyl succinate. The mechanism by which these fuels generate the signals to stimulate secretion is not established. Mitochondrial substrates such as pyruvate and succinate enter the mitochondria in exchange for like charges, or are cotransported with an opposite but equal charge [86]. It should be noted that methyl succinate rather than succinate is added to intact cells because it readily traverses the plasma membrane and is converted to succinate by cytosolic esterases (Fig. 6). The work of MacDonald indicates that malate is the main anion that exits the mitochondria in exchange for succinate [86–89]. This occurs via the dicarboxylate carrier that exchanges succinate for malate with no net anaplerosis. However, net anaplerosis occurs if malate is converted to pyruvate via malic enzyme which also converts nicotinamide adenine dinucleotide phosphate (NADP) to its reduced form NADPH. Pyruvate enters the mitochondria on the pyruvate carrier in exchange for OH− ions and provides both acetyl CoA and anaplerosis at the cost of one proton (for transport) and NADH (to restore malate via malate dehydrogenase). Thus, one signal

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that is generated by the mitochondrial efﬂux of malate is NADPH to increases the cytosolic redox state. It should be noted that glucose also increases cytosolic malate and NADPH production following citrate efﬂux from the mitochondria. Cytosolic citrate forms both acetyl CoA, the precursor of malonyl CoA, and oxaloacetate, which is converted to malate by malate dehydrogenase.

A Role for Cytosolic Redox The cytosolic generation of NADPH by malic enzyme may play a role in the coupling of metabolic to secretory events in nutrient-stimulated insulin release. An important role for redox regulation of signal transduction in cardiac and smooth muscle cells is established [90]. It has long been known that glucose and leucine increase cytosolic redox [91–93]. A recent elegant study by the group of Piston demonstrates the temporal sequence of the rapid increase in cytosolic and mitochondrial NAD(P)H redox state in response to glucose [94]. In intact islets, glucose increases both the reduced to oxidized glutathione ratio (GSH/GSSG) and the tissue content of SH groups. An inhibitor of GSH reductase, 1,3-bis(2-chloroethyl)-1-nitrosourea (BCNU), causes a time-and dose-related, irreversible inhibition of GSH reductase activity [95,96]. This coincides with a fall in both GSH/GSSG ratio and the thiol content of the islets. Pretreatment of islets with BCNU inhibited glucose stimulated insulin secretion implying an important role for either GSH or NADPH. In other cell types, a complex cascade of phosphorylation and dephosphorylation is modulated by redox reactions such that a high redox state of the cytosol increases phosphorylation and degradation of IkB [97,98]. In addition to the GSH system, NADPH also interacts with the thioredoxin (TRX) system [99] (Fig. 7).

Fig. 7. Main cellular redox network controlling oxidation and reduction of thiol residues on proteins via glutathione and thioredoxin systems

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Thioredoxin System Thioredoxin is a pleiotropic cellular component with thiol-mediated redox activity that has been shown to be important in the regulation of many cellular processes in non-beta cells. In diabetic transgenic mice that overexpress TRX in their pancreatic beta cells, the incidence of diabetes is markedly reduced. Streptozotocin induction of diabetes is also attenuated by TRX overexpression in beta cells [100]. A mutant mouse HcB-19 has substantially decreased mRNA expression for TRX-interacting protein (Txnip). Sequencing the critical region revealed a Txnip nonsense mutation in HcB-19 that is absent in its parental strains. Txnip encodes a cytoplasmic protein that binds and inhibits TRX [101].

O2− Generation The mitochondrial electron transport chain is a powerful source of superoxide (O2−). It is composed of four multiprotein complexes: complexes I–IV. Complex I participates in O2− production. Older studies identiﬁed complex III and an unknown component of complex I as the major sites of O2− generation. It has now been shown that the physiologically relevant O2− generation supported by the complex II substrate (succinate) occurs at the ﬂavin mononucleotide group (FMN) of complex I through reverse electron transfer, not at the ubiquinone of complex III [102]. This ﬁnding is compatible with the concept that the major physiologically and pathologically relevant O2−-generating site in mitochondria is limited to the FMN group of complex I. Superoxide dismutase is an essential primary antioxidant enzyme that converts O2− to H2O2. Cytosolic GSH or TRX peroxidase converts H2O2 into water with GSH or TRX as hydrogen donor. Subsequently, GSH or TRX reductase reduces GSSG or oxidized TRX with NADPH as cofactor as described in Fig. 7. NADPH is provided by malic enzyme as malate is converted to pyruvate. The GSH and TRX systems are the major redox buffers in the cell. Peroxidase overexpression alters the cellular contents of GSH, TRX, and reactive oxygen species (ROS) in beta cells [103]. Thus, a metabolically generated increase in NADPH will increase SH content and promote conversion of O2− to water while an increase in O2− will lower SH content or NADPH levels in the course of its metabolism.

A Positive Role of ROS A review of the literature reveals that the role of ROS has been considered primarily with respect to apoptosis and cell death [104–106]. However, it is a normal component of cellular metabolism and we hypothesize that it may

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play another role under non-stressed conditions. Clearly, high levels of O2− trigger dramatic events, but little attention has focused on a transient role during normal metabolism. O2− production is controlled and increases normally under conditions of fuel excess when substrate supply and the mitochondrial membrane potential are high. In addition, the beta cell has unusually low levels of O2− scavenging capacity by superoxide dismutase which would slow the removal of O2− and allow increased ROS production to be sensed, thus providing the potential for serving as an important signal of fuel stimulation. In beta cells, Bindokas and colleagues have recently shown that physiological levels of glucose stimulate ROS production [107]. It should be noted that the intracellular O2− donor tert-butylhydroperoxide in the concentration range of 0.03–30 µM enhances insulin secretion induced by 3-isobutyl-1methylxanthine (IBMX) a non-speciﬁc inhibitor of cAMP and cGMP phosphodiesterases, but mM concentrations of the donor markedly inhibit [108], consistent with a stimulatory role for low concentrations of O2−.

Voltage-Sensing K+-channels MacDonald and Wheeler have documented the presence of redox-sensitive KV channels in pancreatic beta cells [109–113]. These voltage-dependent channels repolarize glucose-stimulated action potentials and are negative regulators of Ca2+ entry and insulin secretion. They did not assess the effect of fuel secretagogues on channel activity in intact beta cells. We hypothesize that the cytosolic generation of NADPH by malic enzyme may target this channel. In the well-studied KATP-dependent signaling pathway of insulin secretion, beta-cell membrane depolarization leads to Ca2+ inﬂux through voltage dependent Ca2+ channels (VDCC). Subsequent repolarization, a requisite for tight control of insulin secretion, is achieved by activation of KCa- and KVchannels. The latter, particularly the KV1 and KV2 families, were identiﬁed recently as predominant contributors to the repolarization of beta-cell membranes, and thus regulate insulin release. Moreover, KV2.1 channels, shown as the major participant, are modulated by the NADPH/NADP ratio. KV channels comprise a diverse superfamily of genes that play important functional roles in excitable cells, including maintenance of plasma membrane potential and repolarization of action potentials.

A Testable Model of Fuel-Induced Insulin Secretion in the Pancreatic Beta Cell To further advance our knowledge, it is essential to put the accumulated metabolic ﬁndings together in a consistent model. The model need not be

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Multiple Roles of High Energy Signals Stimulates ATP/ADP

KATP closure, Respiration, glycolysis kinases

NAD(P)H/NAD(P) Respiration, TRX, KV LC-CoA

Inhibits

Glycolysis, ME

PKC, Lipase, KATP closure, ANT exocytosis

Fig. 8. Opposing functions of the ATP/ADP ratio, redox, and LC-CoA in beta cells. TRX, thioredoxin; ME, malic enzyme; PKC, protein kinase C isoforms; ANT, adenine nucleotide translocase

correct, only testable. As new data accumulate the model should be modiﬁed to reﬂect new data and to continue to provide a testable model for new investigators. The available data provide a number of apparent contradictions (Fig. 8) such as: an elevated ATP/ADP ratio closes the KATP-channels [114] and inhibits respiration [15] while a decrease in the ratio opens the channel [5] and stimulates respiration [15]; LC-CoA has positive effects through stimulation of exocytosis [77] and protein kinase C isoforms [53] and negative effects through opening of KATP-channels [62,63] and stimulating Ca2+-ATPase to lower Ca2+ [58]. To resolve these discrepancies we hypothesize that each of the high-energy signals discussed has dual and alternating roles. Because these signals oscillate, the signal serves a different role at the peaks and at the troughs of the oscillations as indicated in Fig. 8. A hypothetical example of how such a relationship might look is shown in Fig. 9. There are fewer data concerning the relationship between redox and secretion but it has been shown to change with fuel stimulation, oscillate [115,116], and impact secretion when perturbed [92,117]. Among the important metabolic regulatory issues that remain to be resolved are the precise role of redox changes and the mitochondrial/cytosolic compartmentation and pre-secretory time course of adenine nucleotides, redox components, and LC-CoA esters as well as lipids derived from them. Much additional work also needs to be done to identify the redox-interacting proteins and the molecular mechanisms involved in LC-CoA–protein interaction. The proposed model provides an addition, rather than an alternative to the consensus model. This model takes into account the oscillatory behavior of

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Secretion Peaks

LC-CoA, Respiration Peaks

Fig. 9. Hypothetical model of phasing of oscillations in the ATP/ADP ratio, LC-CoA, and insulin secretion. Dashed line identiﬁes a peak in each parameter

metabolism and Ca2+ and suggests a potential role for phasing of oscillations to enhance signal generation. The proposed oscillating interacting metabolic signals that have been identiﬁed are the ATP/ADP ratio, LC-CoA or one of its products, and redox state (the cytosolic NADPH/NADP ratio, ROS, and GSH and TRX redox networks). Acknowledgments. Work from my laboratory on the beta cell has been supported during the past 24 years by NIH grant DK35914. Most of our beta-cell work represents collaborations with Marc Prentki, Jude Deeney, Keith Tornheim, Orian Shirihai, Chris Rhodes, and Gordon Yaney. It has been my privilege to work with an outstanding team of students, post docs, and technicians.

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5. Mechanisms of Beta-Cell Death in Diabetes Marc Y. Donath and Jan A. Ehses

Summary. A decrease in both mass and secretory function of insulin producing beta cells contribute to the pathophysiology of type 1 and type 2 diabetes. In this chapter, we review the evidence that glucose, inﬂammation, dyslipidemia, leptin, autoimmunity, amyloid and some sulfonylureas may contribute to the maladaptation of beta cells. With respect to these causal factors, we focus on IL-1beta, Fas, IRS-2, oxidative stress, NF-kappaB, ER stress, mitochondrial dysfunction, and the KATP-channel as potential mechanisms of action. Interestingly, most of these factors are involved in inﬂammatory processes in addition to playing a role in both the regulation of beta-cell secretory function and cell turnover. To this end, we believe the mechanisms regulating beta-cell proliferation, apoptosis and function are inseparable processes.

Introduction For many years the contribution of a reduction in beta-cell mass to the development of type 2 diabetes was heavily debated. Recently, several studies have convincingly conﬁrmed this hypothesis [1–3], leading to a rapid overemphasis of this etiological factor. Indeed, other mechanisms contributing to the failure of the beta cell to produce enough insulin appear more and more neglected. While we strongly believe that beta-cell destruction is an important etiological factor in the development and progression of type 2 diabetes, in this chapter we will highlight evidence that this is not dissociable from an intrinsic secretory defect. To this end, we will show that pathways regulating beta-cell turnover are also implicated in beta-cell insulin secretory function. It follows that adaptive mechanisms of function and mass share common regulatory pathways and will therefore act in concert. Depending on the Clinic of Endocrinology and Diabetes, University Hospital Zurich, CH-8091 Zurich, Switzerland

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prevailing concentration and the intracellular pathways activated, some factors may be deleterious to beta-cell mass while enhancing beta-cell function, protective to beta-cell mass while inhibiting function, or even protective to beta-cell mass while enhancing function. The failure of the beta cell in type 2 diabetes is akin to a multi-factorial equation, with an overall negative result. Thus, although we will discuss the factors and mechanisms regulating beta-cell mass individually, only in a minority of diabetic patients does one single etiological factor underlie the failure of the beta-cell. In addition to MODYs (maturity onset diabetes in the young), another common example of this is autoimmune-mediated destruction of beta cells in young, lean individuals. However, given that the incidence of type 1 diabetes increases with obesity [4], that insulin resistance is a risk factor for the progression of type 1 diabetes [5], and that approximately 50% of the general population carry the same genetic predisposition [6], this example already implicates multiple etiological factors. Recognition of betacell destruction not only in type 1 but also 2 diabetes led us to recently propose a unifying classiﬁcation of diabetes [7,8].

Glucose and the Interleukin-1beta-FAS–FLIP Pathway: From Adaptation to Failure Glucose is the key physiological regulator of insulin secretion. Therefore, it appears logical that it also regulates the long-term adaptation of insulin production by regulating beta-cell turnover. Indeed, in all species, short-term exposure of beta cells to increasing glucose concentrations induces proliferation in a concentration-dependent manner [9–11]. However, in Psammomys obesus and humans, the proliferative capacity of these cells is suppressed following a prolonged exposure to increased glucose concentrations. With respect to the role of glucose in beta-cell apoptosis, the importance of the genetic background also appears crucial. In rodent islets, increasing glucose from a physiological concentration of 5.5 to 11 mM decreases apoptosis [12]. Further increases above 11 mM will be either pro- or anti-apoptotic depending on the age of the rodent, the culture conditions, e.g. puriﬁed beta cells versus whole islets, or culture on matrix versus in suspension [10,12,13]. The fact that rodent islets survive best at 11 mM glucose is empirically reﬂected by the standard use of medium containing this concentration of glucose for optimal culture conditions. In contrast, in human and Psammomys obesus islets, an increase in glucose from 5.5 to 33 mM induces a linear and much stronger increase in beta-cell apoptosis [10,11,14] (Fig. 1). This difference in glucose sensitivity may explain why in animals genetically predisposed to diabetes, hyperglycemia increases rates of apoptosis, whereas in rats

TUNEL positive β-cells/Islet

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*

20

5.5 mM glucose 11.1 mM glucose 33.3 mM glucose

15 *

10 *

*

5

0 Diabetes-prone

Diabetes-resistant

Rat

Psammomys obesus

Fig. 1. Glucose-induced beta-cell apoptosis in cultured islets of diabetes-prone and -resistant Psammomys obesus and in normal rat islets. Islets were cultured for 9–10 days in 5.5, 11.1 and 33.3 mM glucose. *P < 0.001 relative to islets at 5.5 mM glucose. Adapted from [10], with permission

following 90% partial pancreatectomy, the incidence of beta-cell apoptosis does not change despite increased glucose levels [15,16]. It is probable that such differences also exist between humans with different genetic predispositions. Indeed, although glucose was capable of inducing beta-cell apoptosis in most batches of human islets, which we studied over the last few years, striking variation was observed in the magnitude of this response [11,17–20]. All of this points to a limitation of most studies performed in cell lines, rodent and human islets. Indeed, the above-mentioned differences in susceptibility to glucose-induced beta-cell apoptosis between Psammomys obesus and rat islets highlight the importance of genetic background. Most studies testing agents potentially involved in the failure of the beta cell are performed on islets which do not carry the genetic background predisposing to diabetes. In the future, with respect to testing the effects of various physiological and non-physiological agents on the beta cell, it will be important to see whether human islets isolated from apparently healthy organ donors differ from diabetic islets. Beta-cell mass increases in conditions of increased demand such as in obesity, and its decrease leads to diabetes [21]. This failure to adapt in

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Hyperglycaemia

glucose disposal

IL-1b Long-Term Adaptation Fas Differentiation ↑ Function↑ FLIP

Proliferation ↑ Apoptosis ↓

Fig. 2. Hypothetical model illustrating the consequence of moderate hyperglycemia on beta-cell production of interleukin (IL)-1beta in parallel with insulin secretion. The paracrine effect of IL-1beta induces Fas engagement, which in the presence of FLIP leads to beta-cell proliferation, differentiation and increased function. From [93], with permission

diabetic patients could be explained by the effect of glucose on beta-cell turnover, as detailed above. One potential mechanism by which glucose regulates beta-cell mass can be seen in Fig. 2. According to this hypothesis, long-term adaptation of the beta cells to conditions of increased demand may be triggered by hyperglycemic excursions [9]. These excursions elicit beta-cell production of interleukin (IL)-1beta [20], followed by Fas receptor up-regulation on the pancreatic beta cell [11,22]. In the presence of the caspase-8 inhibitor FLIP, Fas engagement is directed to beta-cell proliferation. However, excessive glucose stimulation can decrease FLIP and induce Fas-receptor death signaling, switching this adaptive pathway towards deleterious signals and eventually to diabetes [17]. In support of this hypothesis, at low concentrations, IL-1beta promotes beta-cell function and survival via the Fas/FLIP pathway [23]. However, IL-1beta is found in increased amounts in the beta cells of type 2 diabetic patients [20], where there is a concomitant induction of Fas [11] and a decrease in FLIP protein expression [17]. In addition to its effect on beta-cell turnover, hyperglycemia is known to impair beta-cell secretory function [7,24–26]. This glucotoxic effect is evident before apoptosis leads to a signiﬁcant decrease in beta-cell mass. This is most striking in vitro, where a 4-day exposure of human islets to elevated glucose concentrations leads to almost complete ablation of beta-cell secretory function although less than 1% of beta cells are apoptotic [20]. Fas receptor engagement in the presence of FLIP may represent one mechanism whereby

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hyperglycemic excursions regulate beta-cell mass. Based on the theme of this chapter, the Fas pathway may therefore also inﬂuence beta-cell secretory function. Studies in our laboratory have recently demonstrated a role for the Fas pathway in regulating insulin production and release [27]. Clearly, the IL-1beta-Fas-FLIP pathway shown in Fig. 1 is not the sole mediator of pancreatic beta-cell adaptation to glucotoxicity [24–26,28,29]. Rather we view this pathway as one example of multiple mechanisms regulating beta-cell mass and function.

Pancreatic Islet Inflammation: A Role in Both Type 1 and Type 2 Diabetes? Classical type 1 diabetes is associated with autoimmunity and pancreatic islet inﬂammation. However, pancreatic islets from type 2 diabetes patients are known to present with amyloid deposits, ﬁbrosis, and increased cell death. All of these hallmarks are associated with an inﬂammatory response. Indeed, given that pancreatic beta-cell mass is now known to decrease in type 2 diabetes, the immune system is most likely associated with the removal of these apoptotic or necrotic endocrine cells. However, prior to massive cell death and immune inﬁltration pancreatic islets may respond to metabolic stress by producing inﬂammatory factors. As mentioned above, the human pancreatic beta cell produces increased IL-1beta in response to glucotoxicity [20]. Various other inﬂammatory factors and nutrients have also been shown to induce an islet inﬂammatory response in vitro [30–32]. In support of this hypothesis, there is now evidence that islet macrophage inﬁltration occurs in the GK rat, the high-energy-diet fed mouse, the db/db mouse, and in human type 2 diabetes before the onset of severe islet cell death [33]. Given that type 2 diabetes is characterized by hyperglycemia, dyslipidemia, and a clinical inﬂammatory state, increased cellular stress may be critical in precipitating islet inﬂammation in vivo. The impact of islet-derived inﬂammatory factors and islet inﬂammation on beta-cell function and mass may be both beneﬁcial and/or deleterious. As mentioned above, low concentrations of IL-1beta promote beta-cell function and survival [34]. Other chronically elevated cytokines and chemokines in obesity and type 2 diabetes, such as IL-6, IL-8, and MCP-1, may also play a role in pancreatic islet adaptation [35]. Indeed, the immune system is classically involved in both adaptation and repair; however, if this response becomes prolonged, it may become deleterious to organ function. In support of this concept, a clinical study using an IL-1 receptor antagonist to treat type 2 diabetes patients improves beta-cell function with no effect on peripheral insulin resistance [36]. All this emerging evidence reinforces the paradigm

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that islet inﬂammation is involved in the regulation of beta-cell function and survival in both type 1 and type 2 diabetes.

Dyslipidemia, Leptin and Circulating Cytokines: Linking Obesity to Beta-Cell Failure Obesity is the main risk factor for the development of diabetes. It is often part of the metabolic syndrome and is accompanied by dyslipidemia, and increased circulating leptin and cytokine levels. All of these factors have been shown to modulate beta-cell function and survival. The inﬂuence of dyslipidemia on the beta cells of an individual will depend on his/her speciﬁc lipid proﬁle. While some free fatty acids (FFA) and lipoproteins have been shown to be pro-apoptotic for the beta cell, others are protective. For example, long-term exposure of beta cells to saturated fatty acids such as palmitate appear highly toxic, whereas monounsaturated fatty acids such as oleate protect against both palmitate and glucose-induced beta-cell apoptosis [18,37]. It is of interest to note that similar toxic effects are also observed in non-beta cells like cardiac cells [38]. These effects are paralleled by effects on beta-cell function (i.e. saturated fatty acids are detrimental, whereas monounsaturated fatty acids are protective). Lipoproteins may also affect beta-cell survival and function in a similar way, whereby VLDL and LDL are pro-apoptotic while HDL is protective [39,40]. These lipotoxic effects may also be inﬂuenced by the prevailing glycemia [41,42]. Leptin was initially identiﬁed as an adipocyte-derived satiety factor. Recently, leptin has also been considered as a pro-inﬂammatory cytokine due to its structural similarity with other cytokines and its receptor-induced signaling pathways [43]. Interestingly, leptin accelerates autoimmune diabetes in the NOD mice [44], providing an additional link between type 1 and type 2 diabetes. Noteworthy, leptin also promotes other autoimmune diseases including inﬂammatory bowel disease [45], multiple sclerosis [46] and rheumatoid arthritis [43]. We have found that leptin, in addition to its established negative effect on insulin secretion [47,48], induces beta-cell apoptosis via increasing release of IL-1beta and decreasing release of the IL-1 receptor antagonist (IL-1Ra) in human islets [19]. Other cytokines released by adipocytes, including tumor necrosis factor (TNF)-alpha and IL-6, may also modulate beta-cell survival, although it is unclear if the amount released into the circulation is sufﬁcient to affect beta cells [6]. Furthermore, it may well be that these cytokines are only effective in the presence of other cytokines. Details regarding how all the above-mentioned factors affect beta-cell secretory function have been reviewed elsewhere [6].

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IRS-2, NF-kappaB, ER Stress, Mitochondrial Dysfunction, and Oxidative Stress: Regulators of Both Beta-Cell Mass and Insulin Sensitivity Several recently published studies point to common responsive elements in the pathways regulating beta-cell turnover and peripheral insulin effects. A striking example is insulin receptor substrate-2 (IRS-2), which promotes beta-cell growth, survival, and insulin secretion, in addition to its wellrecognized role in insulin sensitivity [49,50]. The role of IRS-2 in regulating beta-cell fate has recently been extensively reviewed [50], and is discussed in the chapter by Rhodes. The transcription factor NF-kappaB is another example of such a common regulator. In the beta cell it may induce apoptosis or promote survival, depending on the kinetics and mode of induction [6,51–53]. NF-kappaB is also involved in the etiology of insulin resistance [54,55]. Recent work has provided evidence that NF-kappaB signaling is involved in the low-grade inﬂammation that occurs in the liver of type 2 diabetic models, contributing to insulin resistance [56,57]. Further, this signaling node may inﬂuence peripheral insulin resistance via actions in myeloid cells [57]. Interestingly, non-steroidal anti-inﬂammatory drugs enhance insulin sensitivity and protect the beta cell from apoptosis via inhibition of NF-kappaB [52–55]. Finally, hyperglycemia and FFA induce activation of NF-kappaB [11,58,59], while attenuation of NF-kappaB inhibits glucose stimulated insulin secretion in beta cells [60]. All these ﬁndings support the concept that NF-kappaB may have a key role in causing both insulin resistance and impaired insulin secretion in type 2 diabetes [59]. Due to the high secretory demand it endures, the endoplasmic reticulum (ER) is very well developed and highly active in the beta cell. This also likely increases the susceptibility of these cells to ER stressors. Endoplasmic reticulum stress might produce signals mediating glucose-induced impairment of function and death [61–63]. However, activation of the unfolded protein response in beta cells in response to ER stress may also provide protection to beta cells by attenuating the ability of cytokines such as IL-1beta to signal and activate the expression of downstream targets [64]. Endoplasmic reticulum stress has recently been observed in obesity and linked to insulin resistance [65,66]. Mitochondrial dysfunction has also been proposed as a common feature of both impaired insulin responsiveness of peripheral tissues and defective beta-cell secretory function and survival [67]. The central role of the mitochondria in insulin secretion and insulin sensitivity underlies this hypothesis. Recently, it was shown that PDX-1 regulates insulin secretion via

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mitochondrial effects, while the role of the mitochondrial uncoupling protein (UCP)-2, in insulin secretion is well established [68–70]. It is interesting to note that PDX-1 is also a critical regulator of beta-cell survival [71,72]. Finally, the pathogenic effect of glucose, possibly in concert with FFA, is also mediated by oxidative stress and may not only affect insulin sensitivity, insulin secretion, and survival, but may also play a role in the development of the secondary complications of diabetes [73–75]. These effects are mainly catalyzed by the generation of reactive oxygen species (ROS) and reactive nitrogen species (RNS), which will ultimately activate stress-induced pathways (NF-kappaB, stress kinases, and hexosamines) to manipulate cell fate.

Anti-Diabetic Drugs: Beneficial or Harmful? Understanding that decreased beta-cell mass is an important factor in the pathogenesis of type 2 diabetes raises a concern regarding the application of drugs potentially harmful to the remaining beta cells. Conversely, protection of beta cells from death presents itself as a new therapeutic target. In this context, modulation of the beta-cell ATP-sensitive K+-channel [KATPchannels, octamers composed of four inwardly rectifying K+-channels (Kir 6.2) and four sulfonylurea receptors (SUR1)] appears particularly interesting. Indeed, closure of the KATP-channels by the sulfonylureas tolbutamide and glibenclamide may induce Ca2+-dependent beta-cell apoptosis in rodent and human islets [12,76,77]. This effect was observed only in vitro and not consistently [78]. However, in an important recent clinical study comparing insulin and sulfonylurea treatment of type 2 diabetes, it was shown that treatment with insulin preserved beta-cell function more effectively than glibenclamide [79]. It remains to be established whether it is the beneﬁcial effects of insulin per se, or the possible beta-cell toxicity of glibenclamide that accounts for this observation. Whereas a deterioration of insulin secretion was seen in patients treated with sulfonylureas in the U.K. Prospective Diabetes Study, those treated with insulin were not evaluated in this regard [80]. Given the possible deleterious effect of some sulfonylureas, alternatives to these as well as alternative insulin secretagogues may have to be considered. When applied at the concentration of their respective circulating half-lives in vitro, repaglinide and nateglinide do not appear to have an apoptotic effect on human islets [77]. In contrast to sulfonylureas, KATP-channel openers may exert protective effects on beta cells [81,82]. In 1976, Greenwood and colleagues were the ﬁrst to report an improvement in insulin secretion after administration of diazoxide for 7 days to diabetic subjects [83]. Similar protective effects were observed more recently in patients classiﬁed with type 1 and type 2 diabetes [84,85]. Finally, other anti-diabetic drugs that have

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emerged as protectors of beta cells from apoptotic stimuli include thiazolidindiones, GLP-1 analogues, and last but not least, insulin [52,77,86,87].

Summary and Proposed Integrated View of Beta-Cell Demise in Type 2 Diabetes We have discussed evidence in support of numerous factors that are potentially deleterious to beta cells. Additionally, some individuals may have limited beta-cell mass early in life due to genetic factors predisposing them to diabetes. However, the demise of insulin-producing cells can be compensated for to a certain degree by regeneration. We propose that glucose plays a central role among those factors contributing to beta-cell “burn-out.” While transient, postprandial, hyperglycemic excursions may predominantly induce

Primary modulators: •Hyperglycemia •Dyslipidemia •Cytokines & Leptin •Macrophages

Secondary modulators: •Autoimmunity •Drugs (e. g. sulfonylureas, GLP1, insulin, aspirin, thiazolidindiones)

Local factors: •Predetermined amount of β-cell mass •Sensitivity to pro-apoptotic signals •Regeneration potential of β-cells •Islets derived cytokines (e. g. IL-1β, IL-1Ra, IL-6, TNFα) •Signaling molecules (e. g. Fas, Flip, IRS-2, NF-κB, ER stress, mitochondrial dysfunction, oxidative stress) •Amyloid

Fig. 3. Proposed model for mechanisms regulating beta-cell mass in type 2 diabetes. Before onset of diabetes, insulin resistance may lead to transient postprandial hyperglycemic excursions. Other factors modulating beta-cell mass may include dyslipidemia, cytokines, and macrophages. Genetic predisposition to diabetes may include a predetermined amount of beta-cell mass, as well as differences in the susceptibility to apoptotic signals and in the regenerative potential of the beta cell. Additionally, induction of local inﬂammatory mediators and cell death may activate the acquired immune system. Finally, drugs may protect or harm the beta cell. From [93], with permission

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beta-cell proliferation in insulin resistant individuals, this adaptive mechanism may fail in the long term and be overridden by beta-cell apoptosis. However, it is unlikely that glucotoxicity acts alone and the negative contribution of macrophage inﬁltration, saturated fatty acids, lipoproteins, amyloid [88,89], leptin, and circulating and locally produced cytokines will further “burn-out” the beta cells (see Fig. 3). These factors will induce apoptosis and/or necrosis, which in the presence of pro-inﬂammatory cytokines may activate speciﬁc immunologic phenomena, which ultimately result in autoimmunity [6,90–92]. Finally, evidence now exists that therapeutic agents may inﬂuence, for the good or the bad, the fate of the beta cells.

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62. Harding HP, Ron D (2002) Endoplasmic reticulum stress and the development of diabetes: a review. Diabetes 51 Suppl 3:S455–461 63. Araki E, Oyadomari S, Mori M (2003) Endoplasmic reticulum stress and diabetes mellitus. Intern Med 42:7–14 64. Weber SM, Chambers KT, Bensch KG, Scarim AL, Corbett JA (2004) PPARgamma ligands induce ER stress in pancreatic beta cells: ER stress activation results in attenuation of cytokine signaling. Am J Physiol Endocrinol Metab 287:E1171–1177 65. Ozcan U, Cao Q, Yilmaz E, Lee AH, Iwakoshi NN, Ozdelen E, Tuncman G, Gorgun C, Glimcher LH, Hotamisligil GS (2004) Endoplasmic reticulum stress links obesity, insulin action, and type 2 diabetes. Science 306:457–461 66. Ozcan U, Yilmaz E, Ozcan L, Furuhashi M, Vaillancourt E, Smith RO, Gorgun CZ, Hotamisligil GS (2006) Chemical chaperones reduce ER stress and restore glucose homeostasis in a mouse model of type 2 diabetes. Science 313:1137–1140 67. Lowell BB, Shulman GI (2005) Mitochondrial dysfunction and type 2 diabetes. Science 307:384–387 68. Brissova M, Shiota M, Nicholson WE, Gannon M, Knobel SM, Piston DW, Wright CV, Powers AC (2002) Reduction in pancreatic transcription factor PDX-1 impairs glucosestimulated insulin secretion. J Biol Chem 277:11225–11232 69. Gauthier BR, Brun T, Sarret EJ, Ishihara H, Schaad O, Descombes P, Wollheim CB (2004) Oligonucleotide microarray analysis reveals PDX1 as an essential regulator of mitochondrial metabolism in rat islets. J Biol Chem 279:31121–31130 70. Lameloise N, Muzzin P, Prentki M, Assimacopoulos-Jeannet F (2001) Uncoupling protein 2: a possible link between fatty acid excess and impaired glucose-induced insulin secretion? Diabetes 50:803–809 71. Kulkarni RN, Jhala US, Winnay JN, Krajewski S, Montminy M, Kahn CR (2004) PDX-1 haploinsufﬁciency limits the compensatory islet hyperplasia that occurs in response to insulin resistance. J Clin Invest 114:828–836 72. Johnson JD, Ahmed NT, Luciani DS, Han Z, Tran H, Fujita J, Misler S, Edlund H, Polonsky KS (2003) Increased islet apoptosis in Pdx1+/− mice. J Clin Invest 111: 1147–1160 73. Robertson RP, Harmon J, Tran PO, Poitout V (2004) Beta-cell glucose toxicity, lipotoxicity, and chronic oxidative stress in type 2 diabetes. Diabetes 53 Suppl 1: S119–124 74. Green K, Brand MD, Murphy MP (2004) Prevention of mitochondrial oxidative damage as a therapeutic strategy in diabetes. Diabetes 53 Suppl 1:S110–118 75. Evans JL, Goldﬁne ID, Maddux BA, Grodsky GM (2003) Are oxidative stress-activated signaling pathways mediators of insulin resistance and beta-cell dysfunction? Diabetes 52:1–8 76. Iwakura T, Fujimoto S, Kagimoto S, Inada A, Kubota A, Someya Y, Ihara Y, Yamada Y, Seino Y (2000) Sustained enhancement of Ca(2+) inﬂux by glibenclamide induces apoptosis in RINm5F cells. Biochem Biophys Res Commun 271:422–428 77. Maedler K, Carr RD, Bosco D, Zuellig RA, Berney T, Donath MY (2005) Sulfonylurea induced beta-cell apoptosis in cultured human islets. J Clin Endocrinol Metab 90:501–506 78. Del Guerra S, Marselli L, Lupi R, Boggi U, Mosca F, Benzi L, Del Prato S, Marchetti P (2005) Effects of prolonged in vitro exposure to sulphonylureas on the function and survival of human islets. J Diabetes Complications 19:60–64 79. Alvarsson M, Sundkvist G, Lager I, Henricsson M, Berntorp K, Fernqvist-Forbes E, Steen L, Westermark G, Westermark P, Orn T, Grill V (2003) Beneﬁcial effects of

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6. Ion Channels and Insulin Secretion David A. Jacobson and Louis H. Philipson

Summary. Pancreatic beta cells respond to glucose and other secretagogues with electrical activity. Action potentials are the primary electrical signal of the beta cell and are shaped by the orchestrated ﬂux of ions through various types of ion channels. The expression of a diverse set of ion channels allows dynamic modulation of action potentials in response to multiple input signals. This chapter provides an overview of the ion channels of the beta cell and their role in the regulation of insulin secretion.

Introduction The beta cell is an electrically active cell. This activity is determined by the concerted action of ion channels, transporters and pumps [1–4]. Glucose, incretins, and other molecules stimulate the electrical activity of the beta cell. In the case of glucose, this activity is organized into slow depolarizing waves with a plateau from which action potentials rapidly ﬁre. Quiescent periods separate the glucose-induced slow waves at membrane potentials below the action potential threshold of the beta cell (Fig. 1). The beta-cell action potential is generated primarily by cation ﬂux through ion channels that are modulated by glucose metabolism [4]. The metabolism of glucose generates adenosine triphosphate (ATP) which binds to and inhibits the inward rectiﬁer potassium channel Kir6.2 (a subunit of the ATP-sensitive potassium channel [KATP channel]) reducing its conductance of K+ and causing an increase in the beta-cell membrane voltage [5,6]. The resulting depolarization activates voltage-gated channels including the L-type Ca2+ channel as well as the Na+ channel which leads to the rising phase of the action potential. Continued depolarization activates voltage-gated potassium channels, in particular Department of Medicine, The University of Chicago, 5841 S. Maryland Ave., MC1027, Chicago, IL 60637, USA

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Fig. 1. Electrical activity of a mouse islet treated with 14 mM glucose exhibiting typical depolarizing waves topped with rapidly ﬁring action potentials

Kv2.1, allowing membrane repolarization and the falling phase of the action potential via K+ efﬂux from the cell. Calcium entry additionally regulates Ca2+-activated potassium channels, which affect action potential repolarization and the duration of the slow wave. The timing of action potential ﬁring is regulated by the inﬂuence and extent of repolarizing versus depolarizing ion ﬂux around an activation threshold for voltage-gated channels. Although simple in shape, the action potential is generated by a complex set of coordinated ion movements through several ion channels.

KATP Channel Regulation of the Action Potential Threshold The resting membrane potential of the pancreatic beta cell is set by the activity of the ATP-sensitive potassium channel (KATP channel) [5,6]. The KATP channel is an inward rectiﬁer K+ channel (discussed in Chapter 24) comprised of two subunits, SUR1 and Kir6.2. Its activity sets the membrane potential of the beta cell near −70 mV, determined by the equilibrium potential of K+ (EK). This is due to the activity of the KATP channel allowing inward K+ ﬂow when the cell is hyperpolarized and outward ﬂow when the cell is slightly positive of EK. The degree of activity of K+ efﬂux is minimally limited by the rectiﬁcation of the K+ channel caused by voltage-dependent inhibition of the KATP current by polyamines and Mg2+ [7]. The KATP channel is sensitive to ATP and when the ratio of ATP/ADP (adenosine diphosphate) increases, as with glucose metabolism, ﬂux through the channel is inhibited [5,6]. Channel activity is also sensitive to cyclic AMP and G-protein coupled signaling pathways. ATP inhibition of the KATP channel allows the beta cell to reach the action potential threshold by reducing the outward K+ ﬂow and resulting in an intracellular accumulation of cations. Mutations in the genes encoding SUR1 (ABCC8) and Kir6.2 (KCNJ11) are rare but important causes of both persistent hyperinsulinemia of infancy and neonatal diabetes, and polymorphisms in KCNJ11 have been associated with type 2 diabetes (discussed in Chapter 22). Inhibitors of KATP channels including sulfonylureas and glitinides are used in the treatment of type 2 diabetes and permanent neonatal

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diabetes caused by activating mutations in Kir6.2 and SUR1. Diazoxide activates the KATP channel and is used to treat some forms of hypoglycemia.

Voltage-Gated Calcium Channels and Islet Calcium Oscillations Reduced KATP channel activity resulting from an increase in the ATP/ADP ratio causes the beta cell to slowly depolarize and reach an activation threshold for voltage-dependent calcium channels (VDCC) [8]. Once activated, the VDCCs of the pancreatic beta cell, primarily the L-type Ca2+ channels Cav1.2 and/or Cav1.3 (depending on species), allow inward Ca2+ ﬂux down its electrochemical activity gradient [8–11]. These channels modulate intracellular Ca2+ levels of the beta cell, which oscillate with a pattern that closely follows the glucose-stimulated slow waves of electrical activity. Oscillations in Ca2+ have been linked to the oscillatory pattern of insulin secretion from the islet [12–15]. Although intracellular Ca2+ changes within the beta cell do not alone account for the action potentials, their activity has been shown to play a role in the upstroke of an action potential and affect Ca2+-activated K+ channels, which may inﬂuence the action potential repolarization and/or ﬁring pattern [16]. Pharmacological inhibition of these Ca2+ channels eliminates insulin secretion, so clearly voltage-dependent Ca2+ channels play an essential role in beta-cell electrical activity and insulin secretion [8]. Action potentials of the beta cell are signiﬁcantly inﬂuenced by Ca2+ inﬂux. When Ca2+ entry is blocked with nifedipine or cobalt, the action potential fails to ﬁre even while the threshold for ﬁring is met by glucose-induced KATP channel inhibition (Fig. 2) [17,18]. Correspondingly, promoting Ca2+ entry via activation of L-type channels with the dihydropyridine drug Bay K8644 prolongs the duration of the action potential [18]. The amount of Ca2+ entry also affects the duration of glucose-induced slow waves. When external Ca2+ is reduced to 1 mM, the islet slow waves are extended with more action

Fig. 2. Electrical activity of a mouse islet in 2 mM glucose (no bar), treated with 14.4 mM glucose (black bar), which causes depolarization and rapidly ﬁring action potentials, and with 400 nM nifedipine (gray bar), which inhibits action potential ﬁring

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potentials and reduced repolarization during quiescent periods [17]. However, when the extracellular Ca2+ is elevated to supraphysiological levels between 5 and 10 mM, islets have shorter slow waves with fewer action potentials per wave and longer quiescent periods with greater repolarization between each slow wave [17,19]. These changes directly correlate with the amount of Ca2+ entry into the beta cell during glucose induced depolarization, and thus the Ca2+ levels in the beta cell determine many of the responses to glucose downstream of electrical activity. Typical beta cells contain around 50 secretory granules docked with L-type Ca2+ channels and this interaction is proposed to directly regulate granule fusion and insulin secretion [20,21]. Overexpression of a region of the Ca2+ channel that interacts with the secretory granules (the 11–111 loop) disrupts normal insulin secretion [21]. This region plays an important role in skeletal muscle allowing interaction of the L-type Ca2+ channel of the T-tubule with the sarcoplasmic reticulum ryanodine receptor, an essential interaction for excitation-contraction coupling in these cells [22]. When the L-type Ca2+ channel is activated during T-tubular depolarization, it transmits a movement-based signal to activate the ryanodine receptor releasing sarcoplasmic reticulum Ca2+ stores [23]. As insulin-containing dense core secretory granules contain ryanodine receptors [24], interaction of the L-type channels and ryanodine receptors may also occur in beta cells. The mechanism linking L-type Ca2+ activation to insulin granule exocytosis is complex and plays a critical role in the regulation of insulin secretion [25] (discussed in Chapter 9).

Voltage-Gated Sodium Channels Extracellular Na+ is required for normal insulin secretion from both human and rodent islets [26–32]. Rodent beta-cell action potentials show increased ﬁring frequency and augmented amplitude immediately following Na+ removal implicating an important role for Na+ inﬂux during the action potential [17]. Human and canine beta cells on the other hand require external Na+ for the propagation of glucose-induced action potentials [29]. The exact mechanism of Na+ entry during glucose induced electrical activity of the islet is complex and may involve many channels including voltage-gated Na+ channels. Two voltage-gated Na+ currents have been identiﬁed in rodent beta cells based on their differences in biophysical activation and inactivation [9,32]. One of the Na+ currents is present only when the beta cell is held at membrane potentials below −100 mV before activation with a depolarizing step [9]. As beta cells are at rest near −70 mV this current is most likely predominantly

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inactivated in beta cells. Another Na+ current is activated with depolarization steps from −40 mV to 0 mV when held at −80 mV near the resting membrane potential of the beta cell [9]. As beta-cell action potentials ﬁre at a threshold of about −40 mV, this Na+ channel might play a role in the upstroke of the glucose-induced action potential. Both of the Na+ channels identiﬁed in the rodent beta cell are inhibited by tetrodotoxin (TTX), a toxin speciﬁc for voltage-gated Na+ channels. Interestingly, islet electrical activity is only affected by TTX under low glucose conditions (5–6 mM glucose) whereas TTX has no effect on electrical activity above 10 mM glucose [32]. As the physiological range of blood glucose rarely climbs above 10 mM, activation of voltage-gated Na+ channels during glucose-induced depolarization will inﬂuence the upstroke of the resulting action potential. Indeed, TTX treatment of beta cells undergoing glucose-induced (at 5 mM) action potentials causes a reduction in amplitude of the action potential and broadens its width [32]. Although TTX sensitive voltage-gated Na+ currents have been identiﬁed in the beta cell, other Na+ channels may also combine to modulate glucoseinduced Na+ inﬂux and resulting cellular excitability. Messenger RNA encoding the voltage-gated Na+ channels Nav1.3 and Nav1.7 has been found in beta cells. Nav1.3 has been identiﬁed in human, canine and rodent islets using reverse transcription–polymerase chain reaction (RT-PCR) [33]. In situ hybridization studies conﬁrmed that Nav1.3 mRNA was expressed in human beta cells [33]. The Nav1.7 mRNA was detected using single-cell RT-PCR in a majority of rodent beta cells tested [9]. Future studies will determine which Na+ channel proteins expressed in the pancreatic beta cell are functionally important, but it does seem that their contribution is species-speciﬁc and has greater importance in human than rodent beta cells.

Calcium-Activated Potassium Channels Calcium entry via Ca2+ channels plays many important roles in beta-cell electrical signaling including a role in activating Ca2+ regulated K+ channels. The three main groups of Ca2+-activated K+ channels of the beta cell are large conductance BK channels, small and intermediate conductance channels (SK and IK), and a slowly activated K+ conductance termed Kslow [34–36]. The kinetics of the beta-cell Ca2+-activated K+ currents allow for roles in both action potential repolarization and slow wave duration. Although the molecular identity of Kslow is undeﬁned, both SK and BK channels are expressed in beta cells [36,37]. BK channel transcripts, including multiple splice variants, are highly expressed in the endocrine pancreas [37]. SK1, -2, -3, and -4/IK1 transcripts have been identiﬁed in rodent islets and

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insulinoma cells by RT-PCR. SK2 and SK3 protein have also been found in mouse islets [36]. The activity of both SK and BK channels in various tissues modulate action potential ﬁring and shape, and thus, these channels may also affect the glucose-induced action potentials of the beta cell.

Action Potential Repolarization The contribution of Ca2+-activated K+ channels to the glucose-induced action potential is complex. Inhibitors of SK4/IK1 and BK channels (charybdotoxin, iberiotoxin, and tetraethylammonium [TEA]) have been used to decipher the roles of these two channels during both glucose-induced islet electrical activity and Ca2+ ﬂuctuations [38,39]. Charybdotoxin blocks a signiﬁcant amount of the total voltage-gated outward current of rodent beta cells [39]. Thus, part of the voltage-gated outward K+ current of beta cells is carried through the BK channel, as IK1 channels are not voltage gated. However, the glucoseinduced action potentials of rodent beta cells are not affected by charybdotoxin or low concentrations of TEA [38–40]. It is only when iberiotoxin is used to block BK channels of glucose stimulated beta cells and the primary voltage-gated K+ channels of the beta cell Kv2.1 is also blocked, that a signiﬁcant increase in action potential amplitude appears [41]. The action potential duration and the quiescent period between action potentials are also increased during combined blockade of Kv2.1 and BK channels during glucose stimulation [41]. This indicates that there may be multiple repolarizing K+ channels activated following the action potential upstroke including BK. Small conductance Ca2+-activated K+ channels of the beta cell may also inﬂuence insulin secretion. Apamin, a speciﬁc inhibitor of SK1–3 channels, has no signiﬁcant effect on glucose-induced electrical activity of rat islets [42]. However, glucose-induced Ca2+ ﬂuctuations in mouse islets are increased in amplitude and frequency with apamin [36]. This effect is modulated in part though the SK3 channel, as knockdown of this channel in an inducible transgenic mouse model reduced the Ca2+ changes induced by apamin [36]. Further studies with speciﬁc inhibition of each of individual SK channel subunits will clarify the role of these channels during glucose-induced insulin secretion.

Slow Wave Duration Calcium regulates the duration of the slow wave in part through its regulation of the Ca2+-activated K+ current Kslow. A transient increase in K+ permeability has been shown to precede the termination of the slow wave [43,44]. The current responsible for the termination of the slow wave has been studied in rodent beta cells by applying bursts of depolarizing pulses closely resembling the action potentials of glucose-stimulated beta cells during the slow wave and then recording the resulting current (Fig. 3) [35]. The current induced,

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Fig. 3A–D. Train of action potentials elicits an outward current in beta cells. A Membrane potential oscillations in a beta cell exposed to 10 mM glucose. The glucose concentration was lowered to 5 mM as indicated above the voltage trace. *The ampliﬁer was switched from the current-clamp into the voltage-clamp mode, the membrane potential held at −70 mV, and the command voltage varied as indicated. B The membrane currents elicited by the pulse train (C, top). Note time-dependent decline of outward current. (C, bottom) Change of holding current displayed on an expanded vertical scale. Note gradual development of a holding current. Same experiment as in B. D Membrane potential recording from the same cell as in C before lowering the glucose concentration. In C and D, the vertical line marks the temporal relationship between cessation of stimulation and the onset of rapid repolarization (left) and the onset of rapid depolarization during the subsequent burst (right). The horizontal lines indicate (from top to bottom) the steady state holding current at −40 mV, the plateau potential from which the cell repolarizes upon termination of the burst, and the most negative membrane potential attained between two bursts. Modiﬁed from [35]

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termed Kslow, during this technique is K+ permeable and inactivates over a time course similar to the quiescent period between glucose activated slow waves typical of beta cells (Fig. 3) [35]. Interestingly, the amount of this current activated and the time course of its activation are directly proportional to the amount of electrical activity preceding the recording [35]. Accordingly, the amount of Kslow current closely follows the Ca2+ levels and its inactivation follows a time course similar to the termination of the Ca2+ wave at the end of the slow wave [35]. Kslow currents recorded from beta-TC3 insulinoma cells are increased by stimulating Ca2+ inﬂux with a L-type Ca2+ channel activator [45]. By contrast, inhibition of Ca2+ inﬂux eliminates the Kslow current and removal of extracellular Ca2+ induces a continued slow wave of depolarization [17,30,31,35,45]. The molecular identity of the Kslow current is unknown despite characterization based on its biophysical and pharmacological properties. The current is inhibited by 70% with high concentrations of the K+ channel inhibitor TEA in primary beta cells [35]. Kslow from primary beta cells is insensitive to charybdotoxin and apamin. Thus, the current is most likely not due to BK or SK channel activity [35]. However, the Kslow current from primary beta cells is reduced by UCL 1684, a nonpeptide SK channel blocker, indicating that the current could still be carried by a member of the SK channel family but not sensitive to apamin [46]. UCL 1684 dramatically affects islet glucose-induced electrical activity leading to a shorter duration and reduced repolarization during quiescent periods [46]. This compound also causes changes in the islet Ca2+ handling, increasing internal free Ca2+ concentrations due to greater depolarization [46].

Voltage-Gated Potassium Channels Regulate Action Potential Repolarization One of the primary functions of voltage-gated K+ channels is repolarization of the action potential, but they serve several other functions as well. The voltage-gated K+ channels are comprised of eleven families, of which at least three are found in beta cells (Table 1) [47,48]. The voltage-gated K+ channels were originally named according to homology to three Drosophila genes: Shaker (Kv1.x), Shab (Kv2.x), and Shaw (Kv3.x). Members of Kv1, -2, and -3 families give rise to delayed rectiﬁer-type currents. A fourth family, Shal (Kv4.x) comprises only rapidly activating and inactivating channels (similar to Kv1.4) and are unlikely expressed in beta cells [49]. K+ channel families 5 to 11 include some that do not express functional channels on their own (Kv6–9). Some subunits also form heteromultimers, generating additional current types with combinatorial properties [49,50]. The ability of isoforms to heteromultimerize is usually restricted and family speciﬁc. For example,

Ion Channels and Insulin Secretion Table 1. Expression of Kv channels in pancreatic islets Gene Alias Expression Method KCNA1 KCNA2 KCNA3 KCNA4 KCNA5 KCNA6 KCNA7 KCNA10 KCNB1 KCNB2 KCNC1 KCNC2 KCNC3 KCNC4 KCND1 KCND2 KCND3 KCNF1 KCNG1 KCNG2 KCNG3 KCNG4 KCNS1 KCNS2 KCNS3 KCNH1 KCNH2 KCNH3 KCNH4 KCNH6 KCNH7 KCNH8

Kv1.1 Kv1.2 Kv1.3 Kv1.4 Kv1.5 Kv1.6 Kv1.7 Kv1.10 Kv2.1 Kv2.2 Kv3.1 Kv3.2 Kv3.3 Kv3.4 Kv4.1 Kv4.2 Kv4.3 Kv5.1 Kv6.1 Kv6.2 Kv6.3 Kv6.4 Kv9.1 Kv9.2 Kv9.3 Kv10.1 Kv11.1 Kv12.2 Kv12.3 Kv11.2 Kv11.3 Kv12.1

None None None Beta cell Beta cell Beta cell Islet ND Beta cell Delta cell Alpha cell Beta cell None Delta cell Pancreas Islet Alpha cell None Alpha cell Beta cell None ND ND Islet Beta cell Islet Islet ND ND ND ND ND

99

Ref.

PCR, IB PCR, IB PCR, IB PCR, IB PCR PCR, IB PCR

[60] [60] [60] [60] [56,57] [60] [48]

PCR, IB, IHC PCR, IHC PCR, ISH PCR, ISH, IHC PCR PCR, IHC PCR PCR, IB PCR, IHC PCR PCR, ISH PCR, ISH PCR

[48,60,63] [48] [48] [48,59] [48] [48,94] [48] [66] [94] [48] [48] [48] [48]

PCR, ISH PCR, ISH PCR PCR

[48] [48] [48] [48]

ND, not determined; PCR, polymerase chain reaction; IB, immunoblot; IHC, immunohistochemistry; ISH, in situ hybridization

Kv1.x can combine with other Kv1.x, but not with Kv2.x [49]. Some examples of “promiscuous coupling” of Kv isoforms have been reported, such as between Kv2.1 and Kv9.3 [51,52]. Kv5.x, 6.x, 8.x and 9.x may actually be subunits that are primarily involved in modulating Kv2.x expression or function. The heteromultimers produce channels that attenuate the amplitude of Kv2. x currents. Each modulatory subunit has its own speciﬁc properties on regulation of the functional Kv2 subunits, and they can lead to extensive inhibitions, to large changes in kinetics, and/or to large shifts in the voltage dependencies of the inactivation process.

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Fig. 4A–D. GxTX-1E broadens the beta-cell action potential. A,B Action potentials induced by 10 mmol/l glucose were recorded before (A) and after (B) the addition of 43 nmol/l GxTX-1E. C Representative action potentials from A and B are aligned at the peak to illustrate action potential broadening by GxTX-1E. D Action potentials triggered by injection of positive current (100 pA) before and after the addition of 100 nmol/l GxTX1E. Modiﬁed from [64]

The beta-cell delayed rectiﬁer current activates slowly and inactivates slowly (Fig. 4). Rapidly inactivating Kv currents are usually not seen in primary beta cells. The Kv current is sensitive to TEA in the 5–7 mM range. Several investigators have made an effort to deﬁne the K+ channel gene or genes encoding this current. A summary of all the known voltage-gated K+ channel genes and their expression (or lack thereof) in islets and beta cells is shown in Table 1. Seven different isoforms of the Kv1.x gene family were initially identiﬁed in insulinoma cell lines and whole islets using RT-PCR [53]. These channels were identiﬁed by their sequence similarity to the rat brain K+ channels [54,55]. Kv1.5 (also termed hPCN1, KCNA5) was found to be highly expressed in human insulinomas [56]. Insulinoma cells overexpressing this channel had an ablated glucose-induced Ca2+ response and transgenic mice overexpressing Kv1.5 speciﬁcally in beta cells exhibited impaired glucose tolerance [56]. Inhibiting Kv1 channels with a dominant-negative construct has little to no effect on beta-cell repolarization [57]. Kv3.2 channel RNA and protein has also been identiﬁed in beta cells; however, TEA-sensitive currents do not reﬂect the low sensitivity of the beta-cell Kv-currents to TEA. Kv2.1 channels are the primary beta-cell delayed rectiﬁer current [47,58,59]

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indicated by the voltage-gated kinetics of the current, as well as RT-PCR and Western blot conﬁrmation of its expression speciﬁcally in beta cells [59–61]. Accordingly, most of the voltage-gated K+ current is eliminated in rat beta cells expressing a Kv2 dominant-negative construct [60]. Spider toxins that inhibit Kv2 channels also inhibit a majority of the Kv current of beta cells [62–64]. Thus, Kv2.1 channels appear to play an important role in the action potential repolarization of the beta cell. Speciﬁc inhibition of Kv2.1 with a spider toxin-like peptide, GxTX-1E, during trains of glucose-induced action potentials results in a signiﬁcant increase in the amplitude and the duration of the action potential (Fig. 4) [64]. This change in electrical activity is reﬂected in the Ca2+ ﬂuctuations of the islet, which usually show increased rates in response to inhibition of Kv2.1 during glucose stimulation [63,64]. Similarly, glucose-stimulated insulin secretion is also increased in islets and beta cells when Kv2.1 channels are inhibited [64,65]. Kv2.1 would thus be a therapeutic target if it could be speciﬁcally blocked in the beta cell [47,64,65]. This could allow increased glucoseinduced insulin secretion in patients with type 2 diabetes. However, the Kv2.1 channel is expressed in many other tissues besides the pancreatic beta cell, including the brain, and speciﬁcally blocking Kv2.1 in only the beta cell is a difﬁcult task. Unique inhibitors speciﬁc to Kv2.1 are currently being designed and their ability to amplify glucose-stimulated insulin secretion is being examined [64]. Modulation of beta-cell function through second messengers may also provide beta-cell speciﬁc regulation of Kv2.1 and affect glucose-induced insulin secretion. Glucagon-like peptide 1 (GLP-1) agonists have been shown to reduce Kv2.1 currents [66]. Neuronal Kv2.1 channel activity in the brain is also dynamically modulated by its phosphorylation status in a Ca2+ dependent manner [67]. Kv2.1 activity is also reduced and its inactivation is accelerated by the polyunsaturated fatty acids arachidonate and linoleic acid [68–70]. Hydrolysis of esteriﬁed linoleic acid and arachidonic acid ampliﬁes glucose-induced Ca2+ inﬂux and insulin secretion [70,71]. Thus, dynamic modulation of Kv2.1 channel function by various signaling mechanisms may play an important role in regulating the electrical responses of beta cells to glucose.

TRP Channels of the Beta Cell The transient receptor potential (TRP) cation channel family comprises 28 members in six subfamilies. The pancreatic islets and their associated nerves express members of at least three subfamilies (TRPML, TPPV1, and TRPC). The beta cell expresses TRPM2, TRPM4, TRPM5, TRPC1, TRPC4, TRPC6, and

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Fig. 5A–C. TRPM4 suppression affects insulin secretion. A Effect of TRPM4 protein suppression on insulin secretion under static incubation conditions. Inhibition of TRPM4 by a dominant negative effect signiﬁcantly decreased the response to 10 and 25 mM glucose stimulation compared to control cells. B A signiﬁcant decrease in insulin secretion was also observed in arginine vasopressin stimulated cells. In this experiment, the response to KCl or l-arginine did not differ. Control cells were transfected with reagents without the DeltaN-TRPM4 DNA. Values are mean ± SEM (n = 4 wells/treatment group from three different cell passages; *P < 0.05 compared to same concentration). C Effect of TRPM4 suppression on insulin secretion under perfusion conditions. Inhibition of TRPM4 by a dominant negative effect signiﬁcantly decreased the response to 25 mM glucose stimulation compared to control cells. INS-1 cells were perfused for 10 min with KRB containing 4 mM glucose to obtain a basal level and stimulated with 25 mM glucose for 20 min to induce insulin secretion. At the end, cells were depolarized with 20 mM KCl to test their viability. Control cells were transfected with reagents without the DeltaN-TRPM4 DNA. Modiﬁed from [77]

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TRPV1 [72–78]. These channels pass cations, mainly Na+ and Ca2+, down their respective electrochemical gradients in response to multiple stimuli, thus affecting cellular depolarization. The ﬁrst TRP channel identiﬁed in the beta cell was TRPC1 [72] although TRP-like channel currents had been recorded many years earlier [79,80]. Subsequent studies identiﬁed TRPC4, TRPC6, TRPM4, and TRPM5 in islets and insulinoma cell lines [72–77]. Another TRPM member, TRPM2, has also been identiﬁed in human and mouse beta cells. This channel is activated by hydrogen peroxide and has been implicated in beta-cell apoptosis [74,75,81,82]. Along with the canonical (TRPC) and melastatin (TRPM) channels, a member of the vanilloid family of TRP channels, TRPV1, is expressed in islets and insulinoma cell lines [78]. TRPV1 is also expressed by pancreatic sensory afferent nerves and it may serve several functions in islet biology including regulating beta-cell function directly and indirectly through neural input into the islet [78,83–86]. The TRP channels are likely to be activated during glucose-stimulated insulin secretion and may contribute to the depolarizing Na+ current that leads to activation of voltage-dependent channels as well as Ca2+ entry and intracellular regulation of Ca2+ ﬂux. TRP channels have been reported to affect the electrical activity that occurs during glucose-stimulated insulin secretion. Trivalent molecules that block some TRP channels in heterologous expression systems inhibit glucoseinduced electrical activity and insulin secretion from beta cells. However, they may also block channels other than the TRPs including Ca2+ channels. SKF 96365, a nonspeciﬁc inhibitor for TRP channels, inhibits glucose-induced Ca2+ oscillations in beta cells [73], consistent with a role for TRP channels in glucose-stimulated insulin secretion. Recently, a dominant-negative construct targeting the TRPM4 channel has been shown to reduce Ca2+ ﬂuctuations and insulin release from beta cells (Fig. 5) [77]. TRPM4 channels are sensitive to Ca2+. They have increased trafﬁcking to the membrane and are activated in a Ca2+ dependent manner [77]. The role of TRPM4 in glucosestimulated insulin secretion has yet to be determined. The activity of individual TRP family members during glucose-stimulated electrical activity of the beta cell is an area of intense interest, awaiting development of speciﬁc inhibitors and genetically modiﬁed mouse models.

Chloride Channels Less is known of the role of Cl− channels in beta-cell function compared to other channel types. A voltage-dependent Cl− channel, ClC3, has been localized to the insulin secretory granules and is thought to play an important role in establishing the acidic pH of the granule [87]. The cystic ﬁbrosis

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transmembrane conductance regulator (CFTR), which is involved in the transport of Cl− ions, is also expressed in beta cells. Patients with cystic ﬁbrosis frequently develop cystic-ﬁbrosis-related diabetes mellitus. Whether this is related to the mutations in CFTR that cause cystic ﬁbrosis in the ﬁrst place or is a secondary manifestation of the pancreatic destruction and high caloric load these patients must consume is so far not clear [88].

Ligand-Gated Channels A growing body of literature has established the presence of ATP, glutamate, and gamma-aminobutyric acid (GABA) receptors in insulinoma cells, islets, and isolated beta cells [89–93]. They are likely to be involved in autocrine and paracrine signaling within the islet and may also be involved in regulation of glucagon secretion [89–93]. While speciﬁc isoforms have been identiﬁed in several species, the exact contribution of these channels to islet function is still emerging.

Conclusions In this chapter, we have presented a brief review of the major cation (Ca2+, Na+, and K+) channels of the beta cell. We have also mentioned Cl− channels and ligand-gated channels whose exact role in the beta cell is still to be worked out. The central role of the beta cell in all types of diabetes has increased the importance of understanding how electrical activity induced by glucose and other secretagogues is established and how it is altered in various diabetic states.

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60.

61.

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74. Qian F, Huang P, Ma L, Kuznetsov A, Tamarina N, Philipson LH (2002) TRP genes: candidates for nonselective cation channels and store-operated channels in insulinsecreting cells. Diabetes 51 Suppl 1:S183–S189 75. Inamura K, Sano Y, Mochizuki S, Yokoi H, Miyake A, Nozawa K, Kitada C, Matsushime H, Furuichi K (2003) Response to ADP-ribose by activation of TRPM2 in the CRI-G1 insulinoma cell line. J Membr Biol 191:201–207 76. Prawitt D, Monteilh-Zoller MK, Brixel L, Spangenberg C, Zabel B, Fleig A, Penner R (2003) TRPM5 is a transient Ca2+-activated cation channel responding to rapid changes in [Ca2+]i. Proc Natl Acad Sci USA 100:15166–15171 77. Cheng H, Beck A, Launay P, Gross SA, Stokes AJ, Kinet JP, Fleig A, Penner R (2007) TRPM4 controls insulin secretion in pancreatic beta cells. Cell Calcium 41:51–61 78. Akiba Y, Kato S, Katsube K, Nakamura M, Takeuchi K, Ishii H, Hibi T (2004) Transient receptor potential vanilloid subfamily 1 expressed in pancreatic islet beta cells modulates insulin secretion in rats. Biochem Biophys Res Commun 321:219–225 79. Sturgess NC, Carrington CA, Hales CN, Ashford ML (1987) Calcium and ATP regulate the activity of a non-selective cation channel in a rat insulinoma cell line. Pﬂugers Arch 409:607–615 80. Sturgess NC, Carrington CA, Hales CN, Ashford ML (1987) Nucleotide-sensitive ion channels in human insulin producing tumour cells. Pﬂugers Arch 410:169–172 81. Herson PS, Ashford ML (1997) Activation of a novel non-selective cation channel by alloxan and H2O2 in the rat insulin-secreting cell line CRI-G1. J Physiol 501: 59–66 82. Herson PS, Lee K, Pinnock RD, Hughes J, Ashford ML (1999) Hydrogen peroxide induces intracellular calcium overload by activation of a non-selective cation channel in an insulin-secreting cell line. J Biol Chem 274:833–841 83. Karlsson S, Scheurink AJ, Steffens AB, Ahren B (1994) Involvement of capsaicinsensitive nerves in regulation of insulin secretion and glucose tolerance in conscious mice. Am J Physiol 267:R1071–R1077 84. Carlsson PO, Sandler S, Jansson L (1996) Inﬂuence of the neurotoxin capsaicin on rat pancreatic islets in culture, and on the pancreatic islet blood ﬂow of rats. Eur J Pharmacol 312:75–81 85. Razavi R, Chan Y, Aﬁﬁyan FN, Liu XJ, Wan X, Yantha J, Tsui H, Tang L, Tsai S, Santamaria P, Driver JP, Serreze D, Salter MW, Dosch HM (2006) TRPV1+ sensory neurons control beta cell stress and islet inﬂammation in autoimmune diabetes. Cell 127:1123–1135 86. Gram DX, Ahren B, Nagy I, Olsen UB, Brand CL, Sundler F, Tabanera R, Svendsen O, Carr RD, Santha P, Wierup N, Hansen AJ (2007) Capsaicin-sensitive sensory ﬁbers in the islets of Langerhans contribute to defective insulin secretion in Zucker diabetic rat, an animal model for some aspects of human type 2 diabetes. Eur J Neurosci 25: 213–223 87. Barg S, Huang P, Eliasson L, Nelson DJ, Obermuller S, Rorsman P, Thevenod F, Renstrom E (2001) Priming of insulin granules for exocytosis by granular Cl− uptake and acidiﬁcation. J Cell Sci 114:2145–2154 88. Stalvey MS, Muller C, Schatz DA, Wasserfall CH, Campbell-Thompson ML, Theriaque DW, Flotte TR, Atkinson MA (2006) Cystic ﬁbrosis transmembrane conductance regulator deﬁciency exacerbates islet cell dysfunction after beta-cell injury. Diabetes 55:1939–1945 89. Rorsman P, Berggren PO, Bokvist K, Ericson H, Mohler H, Ostenson CG, Smith PA (1989) Glucose-inhibition of glucagon secretion involves activation of GABAAreceptor chloride channels. Nature 341:233–236

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90. Gonoi T, Mizuno N, Inagaki N, Kuromi H, Seino Y, Miyazaki J, Seino S (1994) Functional neuronal ionotropic glutamate receptors are expressed in the non-neuronal cell line MIN6. J Biol Chem 269:16989–16992 91. Inagaki N, Kuromi H, Gonoi T, Okamoto Y, Ishida H, Seino Y, Kaneko T, Iwanaga T, Seino S (1996) Expression and role of ionotropic glutamate receptors in pancreatic islet cells. FASEB J 9:686–691 92. Wang CZ, Namba N, Gonoi T, Inagaki N, Seino S (1996) Cloning and pharmacological characterization of a fourth P2X receptor subtype widely expressed in brain and peripheral tissues including various endocrine tissues. Biochem Biophys Res Commun 220:196–202 93. Poulsen CR, Bokvist K, Olsen HL, Hoy M, Capito K, Gilon P, Gromada J (1999) Multiple sites of purinergic control of insulin secretion in mouse pancreatic beta-cells. Diabetes 48:2171–2181 94. Gopel SO, Kanno T, Barg S, Rorsman P (2000) Patch-clamp characterisation of somatostatin-secreting cells in intact mouse pancreatic islets. J Physiol 528:497–507

7. Gap Junctions and Insulin Secretion Philippe Klee, Sabine Bavamian, Anne Charollais, Dorothée Caille, José Cancela, Manon Peyrou, and Paolo Meda

Summary. About 30 years ago, pancreatic beta cells were shown to be connected by gap junctions and to exhibit glucose-induced oscillatory electrical activity, two features that were hypothetically linked to insulin secretion. Since then, gap junctions have been shown to be an obligatory feature of beta cells in all species and all physiological conditions studied. They are composed of connexin proteins and allow for beta-cell to beta-cell exchanges of current-carrying ions and other small cytosolic metabolites which synchronize the electrical and metabolic activity of beta cells, and recruit these cells for insulin biosynthesis and release. Together, these effects account for the signiﬁcant contribution of gap junction-dependent signaling to the control of insulin secretion. More recent data suggest that gap junctions, either via the expression of connexin proteins and/or of the intercellular communications that these proteins permit, also signiﬁcantly inﬂuence beta-cell growth, apoptosis, and the resistance of islets to immune attack. The mechanism(s) whereby gap junction signaling exerts these multiple effects is still obscure. Understanding this mechanism is relevant both to our understanding of the physiology of pancreatic islets and to the pathophysiology of beta-cell dysfunction in both type 1 and type 2 diabetes. Furthermore, appropriate expression of gap junctions may be a prerequisite for the engineering of surrogate insulin-producing cells and their proper three-dimensional packaging, which may be important for using these cells as a cell-based therapy for the treatment of diabetic patients. Here, we review the current status of our knowledge in this ﬁeld and its exciting perspectives.

Department of Cell Physiology and Metabolism, University of Geneva, Medical School, 1 rue Michel Servet, 1211 Genève 4, Switzerland

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Introduction Insulin secretion is a multicellular function, which is tightly modulated by a multitude of factors in order to rapidly match the hormonal output with the changing metabolic demand and environmental conditions, particularly with regard to the levels of circulating glucose. To this end, the on and off response of insulin secretion should be rapid, and precisely titrated in quantitative terms through the coordinated function of various numbers of beta cells in different pancreatic islets as well as within each of these microorganisms. The coordinated functioning of pancreatic islets is thought to be achieved by isletto-islet neural and/or hormonal links [1,2]. The coordination of beta cells within each islet is thought to involve signaling molecules diffusing in the intercellular spaces, and simultaneously affecting neighboring beta cells [1,2]. Such molecules are presumably generated within each islet, since isolated islets retain the ability to increase insulin secretion and biosynthesis in response to glucose. However, the ﬁnding that this responsiveness was also retained under conditions which perturb the ﬂux of the endogenous neurotransmitters, hormones, and other signaling molecules that normally circulate in the intercellular spaces of the islets, indicates that this signaling is dispensable for glucose responsiveness, even though it is important for modulating the stimulus-secretion coupling of primary beta cells, as shown by the signiﬁcant changes in threshold, oscillation, and levels of insulin output observed between in vivo and in vitro studies [1,2]. In contrast, disruption of the contacts that beta cells normally establish with each other has been repeatedly shown to result in decreased, if not abolished responsiveness, to glucose, in terms of both biosynthesis and release of insulin [3–7]. These defects are at least partially reversed after cell aggregation [3–7], which led to the hypothesis that surface proteins become functionally activated upon beta-cell to beta-cell contact and are obligatory for proper insulin secretion in response to glucose. Like all other types of epithelial cells, beta cells closely adhere to their neighbors by a variety of cell surface proteins [2,8], many of which are members of multigene families. These proteins selectively interact within restricted domains of the cell membrane to form intercellular junctions. Junctions in turn establish adhesive links between adjacent cells contributing to cell polarity and structural cohesiveness of the islet, and thus allowing for specialized functions of distinct membrane regions. They also establish cell communication allowing for synchronization of different cells via sharing of cytosolic components and provide pathways that transduce signals within and between cells, coupling extracellular changes to intracellular responses and integrating these events throughout the entire islet [2]. Among these

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structures, gap junctions, which were identiﬁed in pancreatic islets more than 30 years ago [8] and implicated in the oscillatory electrical response of beta cells to glucose [9–12], represent a particularly important site of control for the coordination of beta-cell function and for the integrated response of the islets [1,13]. Here, we ﬁrst review gap junction organization and function, and then summarize the available data pointing to the participation of these structures in insulin secretion and other beta-cell functions.

What is a Gap Junction? Gap junctions are membrane domains where channels accumulate (Fig. 1). They allow direct cell-to-cell communication via the intercellular exchange of cytosolic molecules up to 1 kDa in size [13–16]. These channels are formed by membrane-spanning proteins called connexins (Cx). Six Cx molecules oligomerize around a central hydrophilic space to form a tubular structure called a connexon. Because most mammalian cells usually express multiple types of Cxs, homomeric and heteromeric connexons are formed by the oligomerization of six Cxs of the same or of a different type, respectively (Fig. 1). At gap junctions, the connexons of one cell interact in the intercellular space with the connexons of an adjacent cell, to form bi-cellular, cell-to-cell, or junctional channels. Depending on whether the two interacting connexons are of the same or a different composition (Fig. 1), the resulting junctional channel is referred to as homotypic and heterotypic, respectively [13–17]. At gap junctions, the membranes of two adjacent cells are closely apposed (Fig. 1), being separated by an intercellular space reduced to a “gap” of about 2 nm [13–17].

From Genes to Connexins Twenty and 21 genes coding for connexins and distributed over many chromosomes are found in the mouse and the human genome, respectively [14]. Nineteen can be grouped as orthologous pairs (for the nomenclature of the connexin genes see http://www.informatics.jax.org/). The regulation of Cx expression is poorly understood but available evidence indicates that it is mostly at the transcriptional level [14–16]. Biosynthesis of Cx polypeptides appears to proceed like that of most other membrane proteins [15,16], ﬁrst by docking of the nascent polypeptide to a protein channel of the endoplasmic reticulum (ER) membrane. It is during the integration into this membrane that the topographic arrangement of most Cxs is established [15–17]. The Cx proteins have been highly conserved, in both sequence and topography, throughout evolution [17]. They all consist of four membrane-spanning domains, linked by two extracellular and one intracellular loops (Fig. 1), that

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Fig. 1. Gap junctions are specialized domains of the cell membrane. In a schematic view, a gap junction is an area where the membranes of two adjacent cells are closely apposed, deﬁning an intercellular space of only 2–3 nm. By electron microscopy, this “gap” is seen as a central dense line after staining by tannic acid (upper left). At these sites, two adjacent cells concentrate connexons; i.e., hexamers of membrane-spanning proteins which, by freeze-fracture electron microscopy, are seen as uniformly large particles (upper right). At gap junctions, the connexons of one cell pair with the connexons of an adjacent cell to form junctional, cell-to-cell channels. Individual connexons may also diffuse in non-junctional regions of the cell membrane to form hemi-channels. Depending on whether a single or multiple types of connexins oligomerize, homomeric (1 and 2, centre right) and heteromeric connexons (3) form, respectively. In turn, a connexon may pair to a similar or a different type of connexon, resulting in homotypic (2, 3) and heterotypic junctional channels (1), respectively. All connexins span the phospholipid bilayer of the cell membrane, extending from the cytoplasm of one cell to the intercellular space. Their most variable portions, which may be targeted by isoform-speciﬁc antibodies, are located in the former compartment. Thus, immunogold labeling of ultracryosections results in the decoration of the cytoplasmic surface of gap junction plaques (lower left). Bar shown in bottom left panel represents 20 nm in upper right and upper left panels, and 80 nm in bottom left panel

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result in the cytoplasmic localization of both N- and C-terminal regions [14– 17]. The two extracellular loops, which are among the most conserved regions of the proteins, contain three cysteine residues in all but one Cx isoform [14–17]. The four transmembrane domains are also well conserved and form alpha-helical sheets that contribute to the wall of the connexon and line its central hydrophilic space [14–17]. The intracellular loop, on the other hand, is highly variable [14–17]. The N-terminal region is of similar length in all Cxs. However, the C-terminal region differs in length and sequence, facilitating the generation of isoform-speciﬁc antibodies [14–17]. The Cxs have speciﬁc patterns of tissue distribution. Some Cxs are expressed in many cell types, whereas others show a much more limited distribution and are restricted to one or a few cell types [13–17].

From Connexins to Connexons Connexons may assemble anywhere between the ER, the ER–Golgi intermediate compartment and the trans-Golgi network [16], in a way which appears to be Cx-isoform speciﬁc [16,18]. They are probably transported to the plasma membrane by vesicles budding from the ER and Golgi stacks, and fusing with the plasma membrane [16–18], although direct delivery of the proteins by fusion of the ER and plasma membrane has also been proposed [18]. Connexons may be ﬁrst inserted in a non-junctional region of the plasma membrane [16–19], from where they may move to the outer margins of existing gap junction plaques [16–19]. The presence of calcium-dependent adhesion molecules near these plaques [16–19] allows the pairing of the connexons of two adjacent cells, to form junctional channels. Cxs have an unusually short half-life (1–5 h), so that, while newly synthesized connexons are added at the periphery of gap junction plaques, old connexons are removed from their center [16–19]. This turnover is thought to involve the invagination of a vesicle, limited by a gap junction membrane, into one of the two interacting cells, the release in the cytosol of so-called annular gap junctions [20], and the degradation of its connexons via both the lysosomal and proteasomal pathways [16,17].

From Connexons to Junctional Channels Channels made by connexon pairing are unique in at least two respects. First, they are two-cell structures. Second, they not only conduct ions but also larger molecules, including metabolites, nucleotides, morphogens, vitamin cofactors, small peptides, and fragments of nucleic acids [13,16,17,21,22] (Fig. 1). The cell-to-cell exchanges of these molecules via junctional channels is referred to as ionic and metabolic coupling, respectively. Both the conductance and permeability of junctional channels show a high degree of

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selectivity [21–23]. This selectivity is determined both by the Cx composition of the channel and by the funnel-shaped geometry of the hydrophilic channel space, which is longer than that of other channels as it spans two membranes, the intercellular gap, and part of the cytoplasm. The channel has a functional diameter of 2.5–3 nm [14–17]. Channels made by different Cxs can discriminate between closely related molecules such as cGMP and cAMP [21] and some channels favor the passage of nucleotides rather than the cognate base, presumably due to the negative electrical charge that results from phosphorylation of the former molecules [22]. The observation that some channels favor the passage of anions whereas others are almost uniquely selective for cations indicates that the regulation of junctional channels is determined by a combinatorial effect of multiple parameters, including size, charge, and hydrophilicity [23]. Like other membrane channels, the opening of junctional channels is regulated, such that the average open probability of individual junctional channels is about 10%. Experimental gating can be achieved by imposing a voltage difference across the channel, by changing the resting membrane potential, by decreasing the cytosolic pH, or by increasing the concentration of free cytosolic Ca2+[14–17,23]. Whether any of these factors is of physiological relevance remains to be determined, as, individually, each factor is rarely effective in all cell types, and usually needs to be drastically altered, to levels presumably not encountered in vivo. Gap junction channels may also be closed by a variety of drugs, most of which incorporate into the lipid bilayer of the membrane and act as chemical uncouplers [14–17,23]. A few molecules have also been reported to enhance gap junction function, either by increasing the number of junctional channels [22–25] or by improving their gating and/or open probability [26–27].

The Functions of Gap Junctions Gap junctions are thought to serve a large number of functions in both developing and adult tissues, including transmission of beneﬁcial or toxic signals, compensation for enzymatic defects, conferring resistance to toxic conditions, permitting proper embryonic development and morphogenesis, promoting electrical and mechanical synchronization, mediating uptake of nutrients, and controlling cell growth, migration, and secretion [13–17,28]. Initially, most studies of gap junctions were conducted in vitro, using primary cells or transformed cell lines that were grown under conditions favoring or preventing cell contact and junctional communication. Subsequently, a number of the functions which were circumstantially inferred from these studies have been conﬁrmed and/or extended using speciﬁc genetic tools whereby selective Cxs have been altered in genetically modiﬁed mice, including cell-speciﬁc over-expression of selected Cxs and knock-in replacement of

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one Cx species by another [28]. The functional importance of Cxs and junctional communication is also evident from the increasing number of human genetic diseases that have been associated with Cx mutations [14–17,29], as well as the growing list of acquired diseases in which Cx dysfunction is thought to play a central role. It is beyond the scope of this chapter to review all these functions and diseases, and the readers are referred to excellent reviews for a more comprehensive coverage [13–17,28,29]. The following examples were chosen to illustrate some of the functions of Cxs. Gap junctions have been implicated in the prenatal development, morphogenesis, and differentiation of many tissues. The ﬁrst well-documented example of their role in these processes was that of the global deletion of the Gja1 gene, coding for Cx43, which by delaying the migration of the neural crest cells that contribute to cardiac morphogenesis leads to obstructed right ventricular outﬂow, impaired blood supply to the lungs, and perinatal death [30,31]. Even though no similar pathogenic role has as yet been demonstrated in humans, mutations of Cx43 have been reported in patients affected by severe, congenital cardiac malformations [32], and in oculo-dento-digital dysplasia, an autosomal dominant syndrome characterized by craniofacial and limb dysmorphology, spastic paraplegia, and neurodegeneration [33]. Cx mutations result in a large number of genetic disorders, mostly involving the inner ear and the skin. Mutations of several Cxs, most frequently Cx26, causes a variety of nonsyndromic congenital deafness and hyperkeratosis syndromes, highlighting the importance of proper Cx-dependent function and signaling for tissue homeostasis [29]. Gap junctions form the electrical synapses that help synchronize large neuronal and glial populations [14–17]. In turn, this synchronization is thought to be critical for major cognitive processes, including perception, memory, and learning [14]. Of the many Cxs which have been reported in the central nervous system [14–19], only Cx36 has been deﬁnitively shown to form gap junction plaques between neurons [34]. Interestingly, however, Cx36 knock-out mice are viable and do not exhibit major anatomical or functional alterations of the central nervous system [35,36]. They also have normal motor coordination and behavior [35–37]. Presumably, the absence of an obvious neurological phenotype implies compensation for the loss of Cx36. However, mice lacking Cx36 cannot synchronize the oscillatory activity of some neuronal populations [35,36], or coordinate the agonist-induced response of others [38]. These ﬁndings, and the decreased expression of Cx36 observed in models of drug-induced neuronal ﬁring [39], suggest that this Cx may be involved in the development of some forms of epilepsy. Consistent with this idea, a variant of the human Cx36 gene was found to be associated with the juvenile myoclonic form of epilepsy [40]. The mechanism whereby a base change, which does not affect the amino acid sequence of the cognate

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protein, results in disease, remains to be understood. A possibility is that the change affects the stability of Cx36 mRNA, thus resulting in altered levels of the protein [40]. Genetic variation in other Cxs has been implicated in other human diseases [41,42]. It is worth noting that the lack of Cx36 did not desynchronize all types of neurons, notably in regions where cells also express other Cxs [38]. These ﬁndings show that, in spite of the overall similarity in sequence, topography and role in cell-to-cell communication, individual Cxs fulﬁll speciﬁc roles that cannot be compensated by other isoforms. A similar conclusion was reached in a knock-in mouse model, in which the native Cx43 was replaced by the quite different Cx32, under control of the promoter of the former gene, in order for the ectopic protein to be expressed at the same sites and levels as the native one [43–44]. In this case, some tissues showed a normal function, but others did not, indicating that Cx32 could not fully compensate for the loss of Cx43 [43–44], presumably as a consequence of the different permeability, conductance, and gating of the two Cxs [13–22]. Gap junctions are also obligatory features of adult secretory cells of endocrine and exocrine glands, whatever the type of secretory product, the mechanism of its release, or the pattern of regulation by stimuli and inhibitors [1,2,13,45]. A large body of circumstantial evidence indicates that Cxs and/or the junctional coupling that these proteins mediate are not necessary for secretion to occur but are required for its ﬁne regulation, in terms of biosynthesis, storage, and release of many secretory products [1,2,13,45]. Endocrine and exocrine cells have selected distinct, and to a large extent alternative, patterns of Cxs to achieve this regulation [13,45], via differential transcriptional control of various Cxs [13]. Strikingly, when Cx32 was expressed ectopically, either by knock-in of Cx32 at the locus which codes for Cx43 [43] or by transgenic overexpression of the cognate cDNA in a cell type that only expresses Cx36 [46], altered function of the targeted exocrine and endocrine glands was observed [13,43,46], indicating that speciﬁc Cx patterns are required for proper in vivo secretion. An additional regulatory event is indicated by the alterations in the levels of co-expressed Cxs after manipulation of the expression of only one of the genes. Deletion of the basic helix-loophelix transcription factor Mist1 resulted in decreased transcription of the Cx32 gene [47]. It also resulted in decreased levels of Cx26, which were not paralleled by a change of Cx26 mRNA, indicating differential transcriptional, translational, and/or post-translational control of these two Cxs, which are co-expressed by the acinar cells of several exocrine glands [13,45]. Deletion of the Cx32 gene also resulted in altered membrane levels of Cx26 [48], indicating that some stringent, still to be deﬁned mechanism links the expression of different Cxs in various secretory cells [13,45]. At present, no human disease resulting from a severe secretory defect has been directly linked to either Cx dysfunction or mutation.

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Connexins May Exert Effects Independently of Gap Junctions Until recently, most of the functions discussed above and several others that have been attributed to gap junctions were thought to be accounted for by the ability of junctional channels to provide for rapid exchange of cytosolic components between cells [13–22]. However, since the deletion of some Cx gene was not found associated with changes in either the abundance of connexons and/or the extent of junctional coupling, presumably because of compensatory changes of other Cxs, some Cx effects may be dependent on the levels of expression of individual Cxs and/or on the interactions between Cxs and other proteins. Recent data have indeed documented that at least some Cxs can interact with either tight junction-associated proteins, as well as with cytoskeletal proteins [14–17]. Once this set of interactions are deﬁned, it will be possible to revisit which of the effects initially attributed to junctional coupling were indirectly due to other proteins acting downstream of Cxs. Biochemical and biophysical data suggest that some connexons may remain in non-junctional domains of the cell membrane, where they function as “hemi-channels” [15–17,49], i.e., without pairing with a connexon from an adjacent cell (Fig. 1). The uptake by single and clustered cells of membraneimpermeant dyes, as well as the loss from intact cells of membrane impermeant nucleotides, such as adenosine triphosphate (ATP), has been documented and attributed to Cx channels, since these processes can be blocked by uncoupling drugs [49,50]. The hemi-channel differs from a bone ﬁde junctional channel, with respect to the out–in as well as the in–out molecular movements [50]. The opening of an unpaired connexon was expected to cause a rapid and deadly loss of cytosolic molecules. Recent evidence suggests that this is not necessarily the case, because hemi-channels are rare and open only brieﬂy [15–17,49,50]. The existence and relevance of these structures remains to be fully validated in situ. In this regard, the studies using uncoupling drugs are limited by the pleiotropic effects of these drugs and their action on Cx-unrelated membrane channels. Moreover, other channels, such as P2X prurinergic receptors, may account for the ATP release from intact cells [5]. Finally, hemi-channel opening is only seen when exceptionally large potentials steps are established across the cell membrane [50], which are unlikely to be observed in vivo. Future studies should document whether these unusual gating conditions only reﬂect the artiﬁcial environment of the in vitro experiments. Recent studies indicate that Cxs have important but selective effects on the expression of a variety of genes [40,52]. Large-scale analysis of the entire transcriptome of cells expressing different levels of selected Cxs may provide some clues about the mechanism. These observations stress the importance

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of evaluating Cx-dependent signaling whether dependent or independent of gap junction function, and establishing its hierarchy in the complex signaling network that allows individual cells to become integrated in a functionally coherent tissue.

Gap Junctions and Beta Cells Beta-cell gap junctions were identiﬁed more than 30 years ago by electron microscopy [8] (Fig. 2), and their function ﬁrst indirectly suggested by two electrode recordings that documented synchronous waves of electrical activity induced by glucose in different cells of the same islet [9–11]. Further studies showed that these gap junctions typically comprise small numbers of connexons [25] and are associated with tight junctions [8,25,53] (Figs. 2 and 3). The later developments of biochemistry, molecular biology, and genetics, which led to the identiﬁcation of the Cx family, provided an impetus to more directly test the functions of gap junctions in pancreatic islets.

Cx36 Gap Junctions Couple Beta Cells In vivo, beta-cell gap junctions are only comprised of Cx36 [53–56] (Fig. 2), whereas under certain in vitro conditions they may also contain Cx43 [53,57– 59]. Cx36 is a 321 amino acid protein with a long (99 amino acid) cytoplasmic loop containing an unusual stretch of 10 glycine residues, and a short cytoplasmic C-terminal region containing potential recognition sites for casein kinases I and II, cAMP-dependent protein kinase, and calmodulin-dependent protein kinase II [60]. Cx36 does not form functional cell-to-cell channels with other Cxs [23], and the homotypic channels it makes differ from those made by other Cxs by a small unitary conductance (10–15 pS) and minimal voltage sensitivity [61,62]. These features account for the frequent, but small level of electrical coupling that has been documented between islet cell pairs monitored by dual patch-clamp recording [62,63], as well as by the electrophysiological recordings of rodent pancreatic islets in vitro [11,12,64] and in situ [65]. Cx36 channels are fairly permeable to the positively charged exogenous tracer neurobiotin, but less so to negatively charged Lucifer Yellow [66–70], suggesting that they may favor the cell-to-cell transfer of small cationic species. Still, these channels allow for the metabolic coupling of islet cells, as shown by the exchange of negatively charged endogenous molecules, such as phosphorylated glucose metabolites [71] and nucleotides [72]. Most beta cells appear electrically coupled, as indicated by the rhythmical and synchronized bursts of electrical activity as well as by the coordinated Ca2+ oscillations that are observed during glucose-induced insulin secretion throughout intact

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Fig. 2A–C. Pancreatic beta cells are coupled by Cx36 gap junctions. A Electron microscopy shows that the membranes of two adjacent rat beta cells, identiﬁed by the typical ultrastructure of insulin-containing granules, come into contact at a desmosome (d) and a large gap junction (gj). Note that the intercellular space is reduced to a dense, 2 nm-thick gap throughout the latter structure. B Immunoﬂuorescence labeling with speciﬁc antibodies reveals that Cx36 (green) forms minute plaques in the beta-cell-rich region of a mouse pancreatic islet (outlined by the dotted line). Note the paucity of Cx36 at the islet periphery, which is mostly formed by glucagon-, somatostatin-, pancreatic polypeptide-, and ghrelinproducing cells. C The same antibody does not stain the islet from a Cx36-null mouse, validating the speciﬁcity of the labeling shown in B. Bar shown A represents 70 nm A, and 32 µm in B and C

islets [9,10]. Despite this widespread electrical synchronization, the transfer of metabolites and membrane impermeant tracers occurs only between small group of islets cells [66,67,72]. It remains to be demonstrated whether this spatial restriction reﬂects the different sensitivity and speciﬁcity of the electrical and tracer methods, or whether it indicates the existence of distinct metabolic compartments made by different groups of coupled beta cells. The

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Fig. 3A,B. Gap junctions interact with tight junctions in the beta-cell membrane. A Freeze-fracture electron microscopy of an isolated human islet shows the presence within the P-fracture face of a beta-cell membrane of numerous gap junctions (arrowheads). Typically, each junction comprises from a few to a few dozen connexons, is clustered with other gap junctions in restricted domains of the cell membrane, and closely interacts with tight junction ﬁbrils (arrows). B On the opposite, E-fracture face, the same junctions are seen as plaques of pits (arrowheads) and arrays of grooves (arrows), respectively, indicating that their structural proteins span the entire width of each apposed membrane, and seal the intercellular space. Bar: 160 nm

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exclusive expression in beta cells of Cx36 and the capacity of this Cx to only form homotypic channels provides the basis for selectivity in communication, allowing for beta-cell exchanges of signals that should not reach, at least at the same time or rate, other types of islet cells. Such selectivity would be anticipated for proper islet functioning since beta cells and delta cells usually function in parallel, whereas beta cells and alpha cells function antagonistically. The weak dependence of Cx36 channels on transjunctional voltage further suggests that the electrical coupling of beta cells may not be disrupted during the hyperpolarization or the depolarization phases that these cells cyclically undergo during glucose stimulation. Hence, the gap junction pathway allows for beta-cell coupling during both resting and stimulated conditions.

Cx36 Gap Junctions Modulate Insulin Secretion Because the function of gap junctions depends on the functional pairing of at least two cells, a ﬁrst strategy to test the inﬂuence of these structures on insulin secretion was to compare the behavior of single and clustered islet cells. Several studies have documented that single beta cells show increased basal release of insulin, poor or no responsiveness to glucose, decreased basal expression of insulin, decreased proinsulin biosynthesis, and poor or no elevation in free cytosolic Ca2+ after glucose stimulation [3–6,69,73–75]. These defects are at least partially reversible after reaggregation, since beta-cell clustering acutely promotes insulin secretion and biosynthesis induced by glucose stimulation, as well as the handling of free cytosolic Ca2+ [3–6]. Clustering also promotes the recruitment of beta cells that are competent with respect to insulin secretion and biosynthesis, hence decreasing the functional heterogeneity which is observed between single beta cells [3,4,7]. Similar conclusions were reached by exposing isolated islets [73] or intact pancreas [76] to drugs known to block gap junction channels [15–17]. Again, glucoseinduced insulin secretion was found to be altered during islet cell uncoupling, in a way that was rapidly reversible after washout of the uncoupling drug, and which did not signiﬁcantly affect the function of single cells [73]. When Cxs were discovered, novel experiments showed that insulinproducing cell lines that do not release insulin in response to a physiological (i.e., an increase in glucose concentration from 1–2 mM to 5–8 mM) stimulation by glucose do not express Cxs and are uncoupled [68,77]. Cell lines retaining at least some glucose responsiveness expressed Cx36 and were coupled, like primary beta cells [68]. Molecular biology tools allowed investigators to examine the effect of altering Cx expression on islet and beta-cell function. The ﬁrst transgenic mice speciﬁcally tailored for beta-cell studies examined the effect of Cx32, which is not normally expressed in beta cells,

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on islet function [46]. These mice had morphologically normal islets and beta cells. However, they were glucose intolerant since they could not respond normally to glucose concentrations typical of a postprandial glucose load. They also showed only modest insulin release after a supra-physiological glucose challenge [46]. These studies suggest that a large enhancement of beta cell to beta-cell coupling via a Cx which is not normally expressed in the beta cell is deleterious for insulin secretion and glucose homeostasis. Once Cx36 was identiﬁed as the main, if not the sole beta-cell Cx, more physiologically relevant studies were carried out. Thus, Cx36-null mice [78] were found to have morphologically normal pancreatic islets comprised of beta cells lacking gap junctions [70]. These beta cells did not show the normal intercellular synchronization of intracellular Ca2+ transients which is seen during glucose stimulation of beta cells expressing Cx36 and, as a result, do not release insulin in a normal, pulsatile fashion [70]. Furthermore, islets lacking Cx36 showed increased basal release of insulin, explaining why no signiﬁcant increase in hormonal output was observed when the pancreas was challenged by concentrations of glucose found postprandially in the plasma [70]. The excessive basal secretion is consistent with the observation that uncoupled beta cells lacking Cx36 cannot be inhibited by the adjacent cells via the cell-to-cell diffusion of a hyperpolarizing, inhibitory current, an event which is readily observed in situ between the coupled beta cells of control mice [65]. The alterations in insulin secretion observed in the Cx36-null mice are similar to those observed in prediabetic states and type 2 diabetics, indicating that Cx36-dependent signaling may be essential for proper regulation of insulin release [70]. In vitro studies have shown that the secretion control achieved by Cx36 cannot be obtained by other Cxs or E-cadherin [67,79], and that excess Cx36 is as deleterious for insulin secretion as the lack of the Cx [67,77,80]. The results of the in vivo and in vitro studies are consistent, and show that Cx36-dependent signaling involves intra-islet synchronization of the glucose-induced oscillations in intracellular Ca2+ which, in turn, drives the oscillations in insulin output [68,70].

Toward a Model The available data provides compelling evidence for a physiologically relevant role for cell-to-cell communication mediated by Cx36 in the function of beta cells. Glucose stimulation induces secretory and metabolic responses from coupled cells which are stronger than from the same number of single beta cells [3,4]. Thus, coupling between beta cells appears essential to recruit and synchronize the activity of individual cells which, taken individually, are heterogeneous with respect to metabolic and secretory status [13,81]. The identity of the endogenous molecules that permeate connexin channels and

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stimulate or potentiate insulin secretion remains to be established. The most direct evidence presently points to a central role of Ca2+, whose glucoseinduced transients require Cx36 channels to become synchronized across beta-cell populations [68,70]. Why an asynchrony of such waves affects normal beta-cell function remains to be directly tested. Conceivably, irregular Ca2+ oscillations could alter the expression of beta-cell genes or the resistance of beta cells to apoptosis or other events. Preliminary data suggest that this may be the case [40,45,46]. In MIN6 cells, a cell line which responds to glucose and expresses Cx36 as do primary beta cells, a major decrease in Cx36 due to expression of an antisense construct resulted in the loss of insulin secretion [68,79], and led to alterations in gene expression including the induction of hitherto undetected genes [87]. It is plausible that one or more of the proteins encoded by these genes could account for the reduced insulin secretion of uncoupled cells. Recent observations have further raised the possibility that the effects of Cx36 may be mediated via interactions with other membrane proteins, including the claudins and occludin that form tight junctions [53], and the sulphonylurea receptor [82]. Finally, a provocative in vitro study suggests that the levels of Cx36 are downregulated after prolonged exposure to supra-physiological glucose concentrations [83], raising the possibility that loss of Cx36 could account for the desensitization to glucose which is observed under such conditions.

Cx36 and Diabetes The above data strongly support an important role of Cx36 in beta-cell function, raising the possibility that alterations in this protein, in the cell-to-cell communications it establishes, and/or in the molecular signaling it controls, are involved in the pathophysiology of diabetes. At present, this hypothesis has not been investigated in humans. However, experimental data in rodents indicate a plausible role for Cx36 in both type 1 and type 2 diabetes. Type 1 diabetes results from an autoimmune attack directed towards the beta cells. The residual beta-cell mass is insufﬁcient to sustain the insulin demand of the organism. The reasons why some beta cells resist this attack and persist for years within the pancreas of patients remains to be understood [84]. Recent experiments from our laboratory indicate that Cx36 may protect beta cells in vitro as well as in vivo against drugs and cytokines that experimentally reproduce the massive cell death observed at the onset of type 1 diabetes [45,90]. Transgenic mice over-expressing Cx36 were fully protected against the effects of beta-cell cytotoxic drugs (in vivo) and cytokines (in vitro) whereas mice lacking Cx36 were sensitized to their cytotoxicity, and became rapidly hyperglycemic due to loss of most beta cells [45,90]. Thus, Cx channels may signiﬁcantly contribute to the resistance of endocrine cells to

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cytotoxic conditions, either by enhancing their resistance to the damaging conditions and/or by favoring an efﬁcient cell repair mechanism after the insult. Type 2 diabetes is a heterogeneous disorder with a complex pattern of inheritance. Genome-wide scans have already detected linkage of diabetic phenotypes with several loci, including 15q14 [85] where the human Cx36 gene is located [86]. Preliminary data indicate that polymorphisms in the Cx36 gene may be associated with diabetes in a subset of patients, possibly because variation in this gene affect the function or half-life of Cx36, resulting in loss of beta-cell function [40]. In this regard, the alterations observed in rodents lacking Cx36 are similar to the alterations that precede the development of overt diabetes in humans (e.g., loss of insulin oscillations) and, later, characterize the disease (e.g. increased basal release of insulin, failure to increase the insulin output in the presence of postprandial concentrations of glucose).

Cx36 and Beta-Cell Therapy One of the goals of understanding the physiology of beta cells is to develop better, targeted strategies for the treatment and cure of diabetes. One of the most promising therapeutic approaches is the transplantation of surrogate insulin-producing cells that retain the essential features of primary beta cells, including normal stimulus-secretion coupling of glucose-induced insulin release. Efforts are under way in a number of laboratories (see chapters 13,14,15) to identify a source of precursor cells that efﬁciently differentiate into secretory-competent insulin-producing cells. Available studies indicate that such cells function better after assembly into isletlike structures and acquisition of some neuronal traits [88]. Cx36 has been shown to promote the formation of three-dimensional organoids-like structures by beta cells [67], raising the possibility that Cx36 may be important, if not essential, for the production of surrogate beta cells for cell replacement therapy. Interestingly, mouse ES cells lack Cx36 [89] but acquire the protein in parallel with the acquisition of some beta-cell traits [unpublished data]. With regard to type 2 diabetes, it is worth noting that the sulfonylurea glibenclamide increases beta-cell gap junctions and coupling [24,25], raising the possibility that the antidiabetogenic effect of the drug may involve changes in the levels of Cx36 protein, in the function of the resulting channels and/or of Cx36-dependent signaling, in addition to its well-established action on the SUR1-Kir6.2 complex that comprises the ATP-sensitive potassium channel [82]. Thus, Cx36 may be an attractive target for new drugs for the treatment of diabetes.

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Conclusions and Perspectives Most functions of pancreatic beta cells, and notably insulin secretion, are multicellular processes which imply the coordinated functioning of many cells within the pancreatic islet. This intra-islet coordination involves a variety of mechanisms of cell-to-cell communication among which those mediated by the Cx36 channels, which form gap junctions, are prominent. Through these channels, beta cells rapidly exchange cytoplasmic molecules that signal the state of activity of neighboring cells, thus allowing for an integrated, synchronized response of the islet cell community. Recent experiments indicate that the proteins making gap junction channels are required for the ﬁne regulation of the biosynthesis, storage, and release of insulin, particularly in response to glucose stimulation, and that their loss results in pancreatic defects that mimic those observed in type 2 diabetes. Loss of gap junctions also sensitizes beta cells to the cytotoxic conditions thought to prevail at the onset of type 1 diabetes. Other studies suggest that Cxs may affect beta-cell growth and regulate the expression of speciﬁc genes. These ﬁndings raise the intriguing possibility that abnormal Cx36-dependent communication between islet cells may be involved in the pathophysiology of type 1 and type 2 diabetes, in addition to playing an important role in normal islet function. Further studies are required to elucidate the molecular organization of the Cx36dependent pathway, including the identity of the gap junction-permeant signaling molecules and the mechanisms whereby they affect beta-cell function. Acknowledgments. This review is dedicated to Drs. M.J. Parsonage and J.W.A. Turner who described the syndrome that recalled to P.M. the value of a healthy life. The authors thank Mrs. N. Dupont and T. Bollmann for excellent assistance. The Meda team is supported by grants from the Swiss National Science Foundation (310000-109402), the Juvenile Diabetes Research Foundation (1-2005-46 and 1-2007-158), Novo Nordisk, and the Geneva Program for Metabolic Disorders (GeMet).

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74. Giordano E, Bosco D, Cirulli V, Meda P (1991) Repeated glucose stimulation reveals distinct and lasting secretion patterns of individual rat pancreatic B cells. J Clin Invest 87:2178–2185 75. Fernandez J, Valdeolmillos M (2000) Synchronous glucose-dependent [Ca(2+)](i) oscillations in mouse pancreatic islets of Langerhans recorded in vivo. FEBS Lett 477:33–36 76. Bruzzone R, Meda P (1988) The gap junction: a channel for multiple functions? Eur J Clin Invest 18:444–453 77. Vozzi C, Ullrich S, Charollais A, Philippe J, Orci L, Meda P (1995) Adequate connexinmediated coupling is required for proper insulin production. J Cell Biol 131: 1561–1572 78. Guldenagel M, Ammermuller J, Feigenspan A, Teubner B, Degen J, Sohl G, Willecke K, Weiler R (2001) Visual transmission deﬁcits in mice with targeted disruption of the gap junction gene connexin36. J Neurosci 21:6036–6044 79. Calabrese A, Caton D, Meda P (2004) Differentiating the effects of Cx36 and Ecadherin for proper insulin secretion of MIN6 cells. Exp Cell Res 294:379–391 80. Le Gurun S, Martin D, Formenton A, Maeschler P, Caille D, Waeber G, Meda P, Haeﬂiger JA (2003) Connexin36 contributes to control function of insulin-producing cells. J Biol Chem 278:37690–37697 81. Meda P (1996) The role of gap junction membrane channels in secretion and hormonal action. J Bioenerg Biomembr 28:369–377 82. Rocheleau JV, Remedi MS, Granada B, Head WS, Koster JC, Nichols CG, Piston DW (2006) Critical role of gap junction coupled KATP channel activity for regulated insulin secretion. PLoS Biol 4:e26 83. Allagnat F, Martin D, Condorelli DF, Waeber G, Haeﬂiger JA (2005) Glucose represses connexin36 in insulin-secreting cells. J Cell Sci 118:5335–5344 84. Meier JJ, Lin JC, Butler AE, Galasso R, Martinez DS, Butler PC (2006) Direct evidence of attempted beta cell regeneration in an 89-year-old patient with recent-onset type 1 diabetes. Diabetologia 49:1838–1844 85. Mori Y, Otabe S, Dina C, Yasuda K, Populaire C, Lecoeur C, Vatin V, Durand E, Hara K, Okada T, Tobe K, Boutin P, Kadowaki T, Froguel P (2002) Genome-wide search for type 2 diabetes in Japanese affected sib-pairs conﬁrms susceptibility genes on 3q, 15q, and 20q and identiﬁes two new candidate Loci on 7p and 11p. Diabetes 51:1247–1255 86. Belluardo N, Trovato-Salinaro A, Mudo G, Hurd YL, Condorelli DF (1999) Structure, chromosomal localization, and brain expression of human Cx36 gene. J Neurosci Res 57:740–752 87. Mas C, Calabrese A, Caton D, Serre-Beinier V, Meda P (2002) Identiﬁcation of a new gene involved in glucose-stimulated insulin secretion. Diabetologia 45 suppl 2:A85 88. Lumelsky N, Blondel O, Laeng P, Velasco I, Ravin R, McKay R (2001) Differentiation of embryonic stem cells to insulin-secreting structures similar to pancreatic islets. Science 292:1389–1394 89. Huettner JE, Lu A, Qu Y, Wu Y, Kim M, McDonald JW (2006) Gap junctions and connexon hemichannels in human embryonic stem cells. Stem Cells 24:1654–1667 90. Klee P, Charollais A, Caille D, Meda P (2004) Role of connexin proteins in the protection of pancreatic beta cells against cytotoxic attacks. Proceedings Prien post-EASD Meeting: 22

8. Protein Kinase A-Independent Mechanism of cAMP in Insulin Secretion Susumu Seino, Takashi Miki, and Tadao Shibasaki

Summary. Although an increase in the intracellular calcium concentration is the primary signal in the regulation of insulin secretion, other intracellular signals are also important, adenosine 3′,5′-cyclic monophosphate (cAMP) being especially critical. cAMP is well known to potentiate glucose-induced insulin secretion. Until recently, the action of cAMP on insulin secretion was generally thought to be mediated exclusively by the activation of protein kinase A (PKA), which phosphorylates proteins associated with the secretory processes. However, accumulating evidence indicates that the cAMP-binding protein, cAMP-regulated guanine nucleotide exchange factor (cAMP-GEF)/exchange proteins activated directly by cyclic AMP (Epac), participates in a novel, PKA-independent mechanism of cAMP action in insulin secretion. cAMP compartmentation in pancreatic beta cells has been proposed to account for these distinct effects of cAMP signaling in insulin secretion.

Introduction Insulin secretion is regulated by various factors including nutrients, hormones, and neural inputs. Glucose-induced insulin secretion, which is the principal mechanism of insulin secretion, is modulated by intracellular signals such as cAMP and phospholipid-derived molecules, e.g., inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (DAG) [1]. Among these intracellular signals, cAMP is especially important in the potentiation of insulin secretion [1–4]. cAMP itself or cAMP-increasing ligands such as incretins (see below) as well as forskolin strongly potentiate insulin secretion at higher glucose concentrations [5]. It is well known Division of Cellular and Molecular Medicine, Kobe University Graduate School of Medicine, Kobe 650-0017, Japan

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clinically that an oral glucose load elicits greater insulin secretion than an intravenous glucose load when similar blood glucose levels are induced in vivo [6]. This is mainly due to the potentiation of insulin secretion by cAMPincreasing gut-derived factors called incretins such as glucose-dependent insulinotropic polypeptide (GIP) and glucagon-like peptide-1 (GLP-1) [7,8], which are secreted upon glucose ingestion from enteroendocrine K cells and L cells, respectively. It generally has been thought that cAMP exerts its effects on insulin secretion by activating protein kinase A (PKA), which phosphorylates various proteins involved in the secretory process [9]. However, accumulating evidence indicates that cAMP regulates and modulates exocytosis in various secretory cells by PKA-independent as well as by PKA-dependent mechanisms.

The Discovery of cAMP-GEF/Epac, a cAMP Sensor, in Beta Cells cAMP potentiation of insulin secretion is affected by adenylyl cyclase regulators, trimetric G-protein regulators, phosphodiesterase (PDE) inhibitors, and PDE-resistant cAMP analogs that change the cAMP levels in pancreatic islets [2,10–13]. cAMP-increasing ligands potentiate both the early (ﬁrst) phase of a rapid, marked, and transient increase and the late (second) phase at a moderate and sustained increase of glucose-induced insulin secretion [14]. Although cAMP at low glucose concentrations shows little or no effect on insulin secretion [5], a combination of cAMP and high glucose induces potentiation of insulin secretion [15,16]. This indicates that the potentiating effect of cAMP on insulin secretion requires an interaction between cAMP signaling and glucose-induced signaling. It has long been thought that the action of cAMP on insulin secretion is mediated by the activation of PKA and subsequent phosphorylation of various proteins associated with the secretory process of insulin [9,10]. The involvement of PKA in cAMP-mediated cellular functions has been evaluated mostly by using PKA inhibitors. However, the cAMP action that is not blocked by PKA inhibitors has often been ignored. Using capacitance measurements of pancreatic beta cells, cAMP was shown to stimulate exocytosis of insulin granules from a readily releasable pool. This effect was unaffected by PKA inhibition [17], suggesting for the ﬁrst time that cAMP potentiates insulin granule exocytosis in a PKA-independent manner [17]. However, the molecular basis of the PKA-independent action of cAMP in insulin secretion was unknown. During the search for an intracellular molecule that interacts directly with the sulfonylurea receptor SUR1, a regulatory subunit of the pancreatic beta cell ATP-sensitive potassium (KATP) channel, by yeast two-hybrid screening of the insulin-secreting MIN6 cell

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library, a cAMP-binding protein called CAMPS (cAMP sensor) was identiﬁed [18]. CAMPS was found to be a mouse homolog of rat cAMP-GEFII/Epac2, an isoform of cAMP-GEFI/Epac1, which was identiﬁed by sequence database search [19,20]. The interaction of cAMP-GEFII/Epac2 and SUR1 was conﬁrmed by yeast two-hybrid system and in vitro binding experiments [18,21,22]. The accumulating data suggest that cAMP-independent is involved in a cAMP-dependent, PKA-independent process [18,22–26].

General Features of cAMP-GEF/Epac There are two isoforms of cAMP-GEF/Epac, cAMP-GEFI/Epac1 and cAMPGEFII/Epac2 [18–20,27,28]. cAMP-GEFI/Epac1 mRNA is ubiquitously expressed, at high levels in adult tissues including thyroid, kidney, ovary, skeletal muscle, and heart, and at low levels in the brain [18–20]. cAMPGEFII/Epac2 mRNA is predominant in the brain and neuroendocrine and endocrine tissues including pituitary, adrenal, and pancreatic islets. A short form of cAMP-GEFII/Epac2 protein lacking the ﬁrst cAMP-binding domain and the DEP domain is expressed speciﬁcally in the liver [29]. cAMP-GEF/ Epac consists of four functional domains: one or two cAMP-binding domains, a DEP (Dishevelled, Egl-10, and Plekstrin) domain responsible for its localization to the plasma membrane, a REM (Ras exchange motif) domain required for stabilizing GEF (guanine nucleotide exchange factor) activity, and the GEF domain, in which GEF activity toward the Ras-like small GTP-binding proteins, Rap1 and Rap2 takes place [19,20,27]. In addition to SUR1, cAMPGEFII/Epac2 binds to Rab3-interacting molecule 2 (Rim2) [18] and Piccolo [26]. Rim2 is an isoform of the originally identiﬁed Rim (now designated Rim1), a target of the small G-protein Rab3 involved in exocytosis [30]. Piccolo is a CAZ (cytomatrix at the active zone) protein which is thought to deﬁne and organize the site of neurotransmitter release in neurons [31], and forms both homodimers and heterodimers with Rim2 through its C2A domain in a Ca2+-dependent manner [26]. Both Rim and Piccolo have zinc ﬁnger, PDZ, and C2 domains. cAMP-GEF/Epac also interacts with microtubuleassociated proteins [32] and Ras [33]. GTP-bound forms of Rap interact speciﬁcally with effector proteins and activate downstream targets in various cells [34,35]. The molecular mass of cAMP-GEFI/Epac1 and cAMP-GEFII/ Epac2 is about 96.9 and 111.0 kDa, respectively. The dissociation constant of cAMP-GEFI/Epac1 and domain B of cAMP-GEFII/Epac2 for cAMP is in the range of 4.0 and 1.2 µM, respectively [27], while PKA binds cAMP at 5.0– 24.6 nM [36,37]. Accordingly, cAMP-GEF/Epac could be a cAMP sensor in a range at which the PKA activity is fully saturated.

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Studies using X-ray crystallographic and biochemical analyses of cAMPGEF/Epac have proposed a molecular mechanism by which cAMP controls cAMP-GEF/Epac function [38–40]. In the absence of cAMP, a regulatory region containing the cAMP-binding domain interacts directly with a catalytic region containing REM and GEF domains. The interaction of cAMP and cAMP-GEF/Epac is proposed to induce a conformational change of cAMPGEF/Epac through the Val-Leu-Val-Leu-Glu sequence, thereby inducing the GEF activity [41]. Thus, the cAMP-unbound form of cAMP-GEF/Epac inhibits the activity of GEF.

cAMP-GEF/Epac Specific cAMP Analogs To understand cAMP-dependent signaling, cell-permeable cAMP analogs which directly bind to cAMP effectors are a powerful tool for analyzing the downstream signals of cAMP. Amino acid residues in a highly conserved cAMP-binding cassette motif [Gly-Glu-Leu-Ala-Leu-X3–5-Pro-Arg-Ala/ThrAla-Ser/Thr] are required for the interaction with cAMP [38,42]. The binding cassette interacts with the phosphate, ribose ring, and adenine ring of cAMP. A highly conserved glutamate residue in the binding cassette among PKA, cyclic nucleotide gated (CNG) channel, and hyperpolarization-activated cyclic nucleotide-gated (HCN) channel interacts with the 2′-OH of ribose of cAMP. These interactions are important in determining the binding afﬁnity to cAMP [42,43]. However, the glutamate residue is absent in the binding cassette of cAMP-GEFI/Epac1 or domain B of cAMP-GEFII/Epac2. Lysine in domain B of cAMP-GEFII/Epac2, which is a cAMP-GEFII/Epac2-speciﬁc residue, is thought not to be involved in the interaction with 2′-OH of ribose [38]. cAMP analogs modiﬁed at the 6- and 8-positions in adenosine, the 2′position in the ribose ring, and the cyclophosphate ring have been used to examine the biochemical and physiological properties of cAMP-binding proteins (Fig. 1). 8-Bromo-cAMP and 8-pCPT-cAMP bind to both cAMP-GEFI/ Epac1 and the PKA-regulatory subunit with a higher afﬁnity than cAMP [44]. The analogs activate the kinase activity of PKA and the GEF activity of cAMPGEFI/Epac1, respectively [40,45]. Both 6-Bnz-cAMP and the 6-Phe-cAMP, which are modiﬁed at the 6-position, selectively bind to site A of PKA-regulatory substitutes I and II and induce kinase activities. 8-pCPT-2′-O-Me-cAMP, which is modiﬁed at the 2′OH-position of ribose in 8-pCPT-cAMP, exhibits a high binding afﬁnity for cAMP-GEFI/Epac1 (Kd 2.2 µM) and a low binding afﬁnity for the PKA regulatory subunit (Kd 20–30 µM) [44,46]. 8-pCPT-2′-O-Me-cAMP is thought to be a highly selective agonist for cAMP-GEFI/Epac1. In fact, 8-pCPT-2′-O-MecAMP regulates exocytosis in human pancreatic beta cells and rat clonal beta cells [23] and in the nerve terminal of the calyx of Held [47] even in the presence

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Fig. 1A,B. Chemical structure of adenosine 3′,5′-cyclic monophosphate (cAMP) and its analogs. A cAMP. Positions of adenine ring and ribose ring are numbered. Solid line indicates covalent bond. Broken line indicates hydrogen bond. B Epac-selective and protein kinase A (PKA)-selective cAMP analogs. In the cAMP-GEF/Epac-selective analog 8-pCPT2′-O-Me-cAMP, a proton of the hydroxy group at 2′-position of ribose ring and a proton at 8-position of adenine ring are modiﬁed by a methyl (CH3) group and a 4-chlorophenylthio (pCPT) group, respectively. In the PKA-selective analog 6-Bnz-cAMP, a proton of the amino group at 6-position of adenine ring is modiﬁed by a benzoyl (Bnz) group

of the PKA inhibitor H-89 or KT5720. In cardiac myocytes, 8-pCPT-2′-O-MecAMP also selectively induces the accumulation of connexin 43, which forms gap junctions, to cell–cell contacts via the cAMP-GEF/Epac-Rap1 signal, whereas the PKA selective agonist 6-Bnz-cAMP enhances gating function of gap junctions [48]. Thus, 8-pCPT-2′-O-Me-cAMP is a useful tool for discriminating PKA and cAMP-GEF/Epac-mediated effects.

Role of cAMP-GEFII/Epac2 in Insulin Secretion To determine the functional role of cAMP-GEFII/Epac2, not phenochromocytoma PC12 cells were transfected with constructs encoding cAMP-GEFII/ Epac2 cDNA or beta-galactosidase (control) together with human growth

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hormone cDNA to monitor secretion. In control PC12 cells, i.e., PC12 cells transfected with beta-galactosidase, forskolin-potentiated secretion was completely blocked by a PKA inhibitor, H-89, whereas in PC12 cells transfected with cAMP-GEFII/Epac2, forskolin-potentiated secretion was only partially blocked by H-89, indicating that cAMP-GEFII/Epac2 mediates cAMP-dependent, PKA-independent exocytosis [18]. In mouse pancreatic islets, H-89 partially reduced incretin-potentiated insulin secretion, while a combination treatment with H-89 and antisense oligodeoxynucleotides (ODNs) against cAMP-GEFII/Epac2 caused a further reduction of incretin-potentiated insulin secretion [24]. These ﬁndings suggest that potentiation of insulin secretion by incretins is mediated by both PKA-dependent and PKA-independent pathways, the latter involving cAMP-GEFII/Epac2. Treatment of pancreatic islets with antisense ODNs against cAMP-GEFII/ Epac2 reduced both the ﬁrst and second phases of cAMP-potentiated insulin secretion, indicating involvement of cAMP-GEFII/Epac2 in both phases [24].” is to be replaced with “Treatment of pancreatic islets with antisense ODNs against cAMP-GEFII/Epac2 reduced both ﬁrst and second phases of cAMPpotentiated insulin secretion, indicating involvement of cAMP-GEFII/Epac2 in both phases [24]. However, using total internal reﬂection ﬂuorescence microscopy to monitor insulin granule dynamics in isolated pancreatic beta cells, potentiation of the ﬁrst phase but not of the second phase of glucoseinduced exocytosis was found to be markedly reduced in mice lacking cAMPGEFII/Epac2, indicating that cAMP-GEFII/Epac2 is critical for potentiation of the ﬁrst phase of glucose-induced exocytosis [unpublished data, Shibasaki et al.]. The discrepancy between the effect of cAMP-GEFII/Epac2 in the potentiation of insulin secretion in the islet perifusion system and the isolated beta cell system is probably due to the fact that, in contrast to isolated beta cells, pancreatic islets possess three-dimensional structure through cell-cell contacts. Capacitance measurements of beta cells indicated that both GLP-1 and the adenylyl cyclase activator forskolin potentiate exocytosis by enlarging the size of the readily releasable pool and accelerating the mobilization of new granules from the reserve pool [17]. This effect is mediated by both PKAdependent and PKA-independent mechanisms [17]. The PKA-independent (PKA inhibitor Rp-cAMPS-sensitive) component rapidly increased membrane capacitance within 200 ms of membrane depolarization, which triggered Ca2+-dependent exocytosis [25]. The PKA-dependent (Rp-cAMPSsensitive) component occurred mainly during the more than 450 ms of depolarization [25]. This difference between PKA-dependent and PKA-independent components may result from the difference in Kd value, the cAMP concentration at which stimulated exocytosis is half-maximal, because Kd values for these two components are 6 and 29 µM, respectively. Using antisense ODNs against cAMP-GEFII/Epac2 and the Epac-selective cAMP analog 8-pCPT-2′-O-Me-cAMP, the rapid, PKA-independent exocy-

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Potentiation of Insulin secretion Fig. 2. Role of cAMP-GEFII/Epac2 in insulin secretion. cAMP-GEFII/Epac2, by interacting with Rim2 and Piccolo, potentiates insulin secretion. Since cAMP-GEFII/Epac2 activates the small G-protein Rap1, Rap1 probably participates in the exocytotic process of insulin granules. cAMP-GEFII/Epac2 also induces Ca2+ mobilization from the Ca2+ store through the ryanodine receptor (RYR) channel, which may also be involved in the potentiation of insulin secretion [61]

tosis was shown to be mediated by cAMP-GEFII/Epac2 [25]. Considered together, it appears that cAMP-GEFII/Epac2 is responsible for the fast component of cAMP-regulated exocytosis, which requires a high cAMP concentration, while PKA, which is activated at a lower cAMP concentration, promotes the late component. Because cAMP-GEFII/Epac2 interacts with Rim2, cAMP might act through cAMP-GEFII/Epac2 to enable Rim2 to promote the priming of insulin granules (Fig. 2). cAMP-GEFII/Epac2 interacts speciﬁcally with nucleotide-binding fold (NBF)-1 of the SUR1 subunit [18,21] of KATP channels. cAMP-GEFI/Epac1 has also been shown to bind SUR1 [22]. A recent study showed that under conditions in which beta cells are dialyzed with a low concentration of ATP,

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8-pCPT-2′-O-Me-cAMP inhibits KATP channel activity [22]. In a model proposed by Kang and colleagues [22], SUR1 recruits cAMP-GEFII/Epac2 to the plasma membrane where cAMP-GEFII/Epac2 mediates the cAMP-dependent activation of Rap GTPase activity. Rap stimulates PLC-epsilon [49], a phospholipase that catalyzes hydrolysis of phosphatidylinositol 4,5-bisphosphate (PIP2). The ability of cAMP-GEFII/Epac2 to hydrolyze PIP2 may explain the inhibitory action of cAMP at KATP channels. Interaction between cAMPGEFII/Epac2 and SUR1 is not inhibited in the presence of high ATP concentrations [50], suggesting that the ATP concentration does not affect interaction between cAMP-GEFII/Epac2 and SUR1. Because the binding of cAMP to cAMP-GEFII/Epac2 induces its conformational change [51], cAMP-GEFII/ Epac2 could dissociate from the SUR1-cAMP-GEFII/Epac2 complex upon cAMP stimulation. Incretin-induced, PKA-independent insulin secretion is impaired in SUR1 knockout islets [52,53] and early PKA-independent exocytosis is lost in SUR1 knockout beta cells [25], suggesting that the interaction between SUR1 and cAMP-GEFII/Epac2 may be necessary for cAMPdependent, PKA-independent insulin secretion. It has also been proposed that a putative SUR on insulin granules (gSUR) couples with the chloride (Cl−) channel (ClC-3) and interacts with cAMP-GEFII/Epac2 [29]. Opening of the ClC-3 channels allows Cl− entry into the insulin granules, thereby promoting granular acidiﬁcation which is required for priming of the secretory granules and reﬁlling the readily releasable pool of granules [54]. It has been hypothesized that the binding of cAMP to cAMP-GEFII/Epac2, which interacts with gSUR, induces opening of the ClC-3 channels, thereby facilitating the priming step of granule exocytosis [25]. Mobilization of Ca2+ from intracellular Ca2+ stores by activation of cAMP signaling also contributes to insulin secretion. For example, GLP-1 mobilizes Ca2+ from the endoplasmic reticulum in pancreatic beta cells [55] through IP3 receptors and/or ryanodine-sensitive receptors [56]. Accumulating evidence indicates that cAMP mediates Ca2+ mobilization from the ryanodine-sensitive Ca2+ store independently of PKA and that cAMP-GEFII/Epac2 is involved in this process [23,58,59]. 8-pCPT-2′-O-Me-cAMP has been shown to mobilize Ca2+ from intracellular Ca2+ stores [23,60], indicating PKA-independent, cAMP-GEFII/Epac2-mediated Ca2+ mobilization. Several possible mechanisms are proposed [61]: (1) cAMP-GEFII/Epac2 interacts with intracellular cellular Ca2+ release channels (IP3 receptors and ryanodine receptors), thereby promoting their opening in response to Ca2+ or Ca2+-mobilizing second messengers; (2) cAMP-GEFII/Epac2 acts through Rap and ERK to promote the PKA-independent phosphorylation of these channels, thereby increasing their sensitivity to Ca2+ or Ca2+-mobilizing second messengers; and (3) cAMPGEFII/Epac2 acts through Rap to stimulate PLC-epsilon, thereby hydrolyzing PIP2 and generating IP3.

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cAMP Compartmentation in Beta Cells cAMP signals are now known to be localized in distinct microdomains or functional compartments (compartmentation) within a cell, whereby they mediate distinct physiological effects (see [62–64] for recent reviews). cAMP compartmentation was originally proposed in cardiac myocytes, as prostaglandin E1 and the beta-adrenergic receptor agonist, isoprenaline, both elevate cAMP and activate PKA but induce different effects on activation of glycogen phosphorylase [65]. Using whole-cell patch-clamp recordings simultaneously at two different sites in single isolated cardiac myocytes of the frog, it was shown that local application of the beta-adrenergic agonist preferentially stimulated local L-type voltage-dependent calcium channel (VDCC) activities but not distant ones, while local activation of adenylyl cyclase by forskolin resulted in VDCC currents anywhere in the cell [66]. In addition, it was shown that local inhibition of PDE activity strongly reduced nonuniformity of the beta-adrenergic response [67]. These studies support the notion that cAMP effects are compartmented in cardiac myocytes. G-protein coupled receptors, Gs-proteins, adenylyl cyclase isoforms, PKA isoforms, PDE isoforms, phosphoprotein phosphatases, A-kinase anchoring protein (AKAP) isoforms, PKA substrates, and possibly other unidentiﬁed factors all contribute to cAMP compartmentation [68–70]. Thus, cAMP compartmentation may underlie the variety of cAMP-regulated physiological responses seen upon stimulation by various cAMP-increasing ligands [71–73]. cAMP-GEF/Epac might be localized in cAMP compartments distinct from those containing PKA. Because cAMP-GEF/Epac has a much lower afﬁnity for cAMP than for PKA [27,36,37], much higher concentrations of cAMP would be required in a cAMP compartment in which cAMP-GEF/Epacmediated signaling occurs than in a cAMP compartment in which PKAmediated signaling takes place.

Conclusion cAMP signaling in insulin secretion has been well studied but the effects of PKA in cAMP signaling have been investigated mostly by pharmacological approaches using various PKA inhibitors, and the cAMP action not blocked by PKA inhibitors was not investigated in detail. It has now become clear that cAMP-GEFII/Epac2 is a target of cAMP in insulin secretion. Recently, new tools such as FRET-based probes and cAMP effector-speciﬁc agonists have been developed for the analysis of cAMP, PKA, and cAMP-GEF/Epac signals, so the mechanism of cAMP-regulated insulin secretion must be re-examined. Investigation of cAMP compartmentation may clarify the integrative process

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of PKA-dependent and PKA-independent signaling in cAMP-regulated insulin secretion. Acknowledgments. The studies from our laboratory that are cited in this chapter were supported by the Ministry of Education, Culture, Sports, Science, and Technology. We thank Ms. Grace Honkawa for assistance in preparing the manuscript.

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9. Regulation of Insulin Granule Exocytosis Erik Renström* and Patrik Rorsman†

Summary. Insulin is stored in secretory granules within the pancreatic beta cell. The release of insulin requires the fusion of the secretory granule with the plasma membrane and the discharge of the granule contents into the extracellular space. Insulin secretion follows a characteristic biphasic time course consisting of a rapid but transient 1st phase followed by a slowly developing and sustained 2nd phase. Because type 2 diabetes involves defects of insulin secretion, manifested as a loss of 1st phase and a reduction of the 2nd phase, it is important to understand the cellular mechanisms underlying biphasic insulin secretion. Here we describe how glucose, via electrical activity, triggers insulin secretion. With this background, we consider the molecular machinery involved in the exocytosis of insulin, the possibility that Ca2+-inﬂux through different Ca2+ channels underlies phasic insulin secretion, how Ca2+ is sensed by the beta-cell granules, the maintenance of the pool of release-competent granules by intracellular granule trafﬁcking and glucose metabolism, the existence of two parallel pathways of exocytosis in the beta cell, and ﬁnally the evidence suggesting that exocytosis is not an all-or-none event and that signiﬁcant regulation of beta-cell secretion occurs at the levels of the fusion pore (the connection between the granule interior and the extracellular space).

Introduction Insulin is the body’s only blood glucose-lowering hormone and is secreted by the pancreatic beta cells of the islets of Langerhans. It is processed to its biologically active form and stored, pending its release, in secretory granules. *Lund University Diabetes Centre/Department of Clinical Sciences, UMAS Entrance 72, CRC 91-11, SE20502 Malmö, Sweden † Oxford Centre for Diabetes, Endocrinology and Metabolism, University of Oxford, Oxford OX3 7LJ, UK

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Every beta cell contains ∼10000 secretory granules [1,2], each containing up to 8 fg insulin (∼1 million insulin molecules) [3]. The latter amount corresponds to an intragranular insulin concentration of ∼100 mM. In man and in most other species, insulin is stored in crystalline form as a Zn2–insulin6 complex. The high insulin concentration in beta cells is also illustrated by the fact that as much as 5–10% of the protein content per beta cell is insulin. During the last ten years, a number of electrophysiological and optical techniques have been developed that, in addition to allowing beta-cell exocytosis to be monitored at high resolution, also enabled detailed studies of the events that precede granule fusion as well as the actual fusion event itself. Here we discuss some of the insights that have been gained by application of these techniques to the beta-cell. Our focus is on what has been gleaned by electrophysiological measurements. Other chapters in this volume by Nagamatsu (chapter 10) and Takahashi (chapter 11) more speciﬁcally consider data generated by optical approaches.

Electrical Activity Couples Elevation of the Blood Glucose Concentration to Insulin Secretion The beta cell is electrically excitable and uses changes in membrane potential to couple variation in the blood glucose concentration to stimulation or inhibition of insulin secretion. At glucose concentrations <5 mM, the beta cell is hyperpolarized and no electrical activity occurs. In freshly isolated mouse islets,1 application of glucose at concentrations ≥7 mM leads to the induction of bursts of action potentials (active phase) separated by repolarized electrically silent intervals [4,5]. The periods of electrical activity are accompanied by changes in the cytoplasmic Ca2+-concentration [6], which in turn drive pulsatile insulin secretion [7]. In the intact islet, every beta cell has the capacity of responding in a graded fashion to glucose and as the concentration of glucose is elevated, there is a progressive increase in the fraction of the active phase at the expense of the silent intervals. Finally, at glucose concentrations beyond 20 mM, uninterrupted action potential ﬁring occurs. The ability of glucose to elicit electrical activity is referred to as its “triggering action.” Glucose also exerts an “amplifying” action on secretion which is exerted at a level distal to the elevation of intracellular Ca2+ [Ca2+]i [8]. However, glucose is unable to elicit insulin release if electrical activity and the accompanying Ca2+-inﬂux are prevented. Thus, beta-cell electrical activity

Most studies of beta-cell electrophysiology have been conducted on mouse islets or isolated mouse beta cells. Unless otherwise indicated, the description here therefore refers to the situation in mouse beta cells. 1

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is a key element in the series of steps culminating in glucose-induced insulin secretion. The glucose-evoked changes in membrane potential are orchestrated by the concerted activity of a large number of different types (perhaps 10–20) of ion channels, each present in hundreds to thousands of copies [5]. A fuller account of beta-cell ion channels is provided elsewhere in this volume (chapter 6 by Phillipson) and here we concentrate on aspects of immediate relevance to insulin secretion and beta-cell exocytosis. Adenosine triphosphate (ATP)-regulated K+ channels (KATP channels) constitute the glucose-regulated resting conductance of the beta cell. The regulation of these channels and their signiﬁcance for glucose-induced electrical activity and secretion are reviewed elsewhere in this volume (see chapter 20 by Ashcroft). In addition, there is some evidence implicating KATP channels in the generation of the bursts. Ca2+-entry, via activation of Ca2+-removal processes (including the Ca2+-ATPase), has been postulated to result in a fall in the cytosolic ATP/adenosine diphosphate ADP)-ratio that in turn produces a transient increase in KATP channel activity [9]. It has been proposed that the higher the glucose concentration, the greater the capacity of the beta cell to cope with the increased ATP-consumption resulting from increased Ca2+pumping without a reduction in the ATP/ADP-ratio large enough to open the KATP channels. This scenario explains the observation that even though the KATP channels are completely blocked at low glucose concentrations (5–8 mM), electrical activity increases over a much wider range of concentrations as well as accounts for the ability of the KATP channel blocker tolbutamide to convert oscillatory electrical activity into continuous action potential ﬁring. In addition to this indirect effect of electrical activity and Ca2+-entry on the resting K+-conductance, Ca2+ also directly activates small-conductance K+ channels [10,11]. When the KATP channels are fully closed, the beta cell depolarizes and action potentials are initiated. In mouse beta cells, the action potential is dependent on the opening of voltage-gated Ca2+ channels and the resulting increase in cytoplasmic Ca2+-concentration then triggers exocytosis of the insulin granules [12,13]. Pancreatic beta cells contain at least three different types of voltage-gated Ca2+ channel: L, R and P/Q [14,15]. [Ca2+]I oscillations and action potential ﬁring persists following knockout of either R- or L-type Ca2+ channels [14,16] indicating that one of the channels is sufﬁcient to maintain these processes. In rat [17] and human beta cells [18], but not in mouse beta cells, tetrodotoxin (TTX)-sensitive voltage-gated Na+ channels also contribute to the upstroke of the action potential. Glucose-induced insulin secretion from human islets is accordingly strongly inhibited by TTX, whereas this toxin has no effect on hormone release from mouse islets. The downstroke of the beta-cell action potential results from the activation of voltage-gated

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K+ channels. Beta cells express multiple types of voltage-gated K+ channel but there is growing consensus that Kv2.1 is of particular importance [19]. Blockade of voltage-gated K+ channels produces a concentration-dependent stimulation of glucose-induced insulin secretion via a lengthening of the action potential duration and the resultant increase in Ca2+-entry. Both Ca2+ and K+ channels have the capacity of interacting with the secretory apparatus via their capacity to associate with the SNARE-proteins [20,21]. Whereas the signiﬁcance of this association is obvious in the case of the Ca2+ channels (to direct Ca2+-entry to the release sites), this is not immediately evident in the case of the K+ channels. In addition to the above channels, beta cells also express channels regulated by hormones and neurotransmitters. A K+ channel controlled by an inhibitory G protein [22], probably a member of the GIRK family, is activated by inhibitory agonists like galanin, somatostatin, and adrenalin. Opening of these channels repolarizes the beta cell and thus inhibits beta-cell electrical activity and secretion. Importantly, none of these compounds produce complete inhibition of electrical activity [23–26]. Conversely, acetylcholine (ACh) stimulates insulin secretion. This effect is exerted via activation of cation-permeable channels linked to muscarinic M3-receptors [27]. Opening of these channels results in membrane depolarization, increased electrical activity, and stimulation of insulin secretion. However, as discussed below, the effects on electrical activity only account for part of the inhibitory/stimulatory effects of these agonists and modulation of secretion at a late stage is quantitatively signiﬁcant.

Observed Rates of Insulin Secretion are Low Compared with Beta-Cell Insulin Content Insulin secretion evoked by a number of stimuli exhibits a characteristic biphasic pattern that consists of a transient ﬁrst phase followed by a sustained second phase during which secretion continues at a somewhat lower rate but is still enhanced with respect to the pre-stimulatory control (Fig. 1). Although it is often claimed that the second phase is less prominent in the mouse than in the rat, this may simply be a consequence of the plasma glucose concentration in the mouse being higher than that of the rat (8–9 mM vs. 4–5 mM) [14,28]. Whereas insulin secretion in response to 10 mM glucose in mouse islets is essentially monophasic and consists almost exclusively of 1st phase release, 2nd phase insulin secretion becomes prominent at higher glucose concentrations (≥16 mM) (Fig. 1). In man, insulin secretion evoked when plasma glucose is increased by ∼7 mM above basal, 1st phase insulin secretion peaks at 1.4 nmol/min and lasts ∼10 min whereas 2nd phase release amounts to ∼0.4 nmol/min [29].

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dCdt 400 fF/s 2s Fig. 1A–D. Biphasic stimulation of insulin secretion. A Biphasic stimulation of insulin secretion in a rat pancreas in response to 16.7 mM glucose. B,C Insulin secretion observed in the perfused mouse pancreas when glucose was increased to 10 mM (B) or 17 mM (C). Data in C are from [16]. D Biphasic increase in cell capacitance (∆Cm) in response to a step increase in [Ca2+]i from a basal concentration of 0.1 µM to a peak a new plateau of 20 µM. The lower trace shows the time derivative (dc/dt) of a function ﬁtted to the time course of capacitance increase to illustrate the kinetics of exocytosis. Data from [2]

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Given that a human beta-cell granule contains 1.6 amol of insulin [3], it can be estimated that 1st and 2nd phase secretion results from the release of close to <109 granules/min and 0.25 × 109 granules/min. These formidable rates should be considered against the fact that there are 1 million islets per pancreas, each containing ∼103 beta cells, therefore the rates of glucose-induced secretion quoted above correspond to ≤1 granule per beta cell per minute. The number of granules per human beta cell has not been determined but electron micrographs suggest that the ultrastructure of human and mouse beta cells include the number of granules as well as the intracellular distribution of granules. Assuming that a human beta cell contains 104 granules and that 20% of these are docked or nearly docked (as in the mouse [2]), the nearmembrane granules that we wake up with in the morning should be sufﬁcient to take to us through the day In fact, the granules are so plentiful and the rate of release so low that it seems almost inconceivable that diabetes could develop solely as a consequence of insulin deﬁciency.

Molecular Machinery of Exocytosis in Beta Cells Regulated exocytosis of insulin involves a multitude of events starting with the synthesis of the preproinsulin molecule in the endoplasmic reticulum, followed by budding of immature insulin granules from the trans-Golgi network, insulin granule maturation, and ﬁnally fusion of release-competent dense-core insulin granules with the plasma membrane (Fig. 2A). A large number of proteins have been described that control the secretory pathway, but the key molecules controlling membrane fusion are the SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) proteins. In man, the superfamily of SNARE proteins consists of 36 small proteins that share the conserved SNARE motif. The combination of four appropriate SNARE motifs results in the spontaneous formation of extremely stable helical core complexes (Fig. 2B). Each SNARE motif contributes one residue to the highly conserved central part of the core complex that contains four amino acids: three glutamine (Q) and one arginine (R) residues. In the beta cell, as in other secretory cells, the fusion of the dense-core insulin granules with the plasma membrane involves the assembly of a complex between the granule membrane R-SNARE VAMP-2 and the integral plasma membrane Qc-SNARE syntaxin-1a, as well as the membrane-associated Qa-Qb SNARE SNAP-25. A large number of accessory factors interact with SNAREs and inﬂuence the assembly of the core complex [30]. One example is tomosyn-1, which is expressed in beta cells and contains an R-SNARE-motif that can replace VAMP2 in the assembly of a heterotrimeric complex with syntaxin-1 and SNAP25 [31] (Fig. 2C). Knockdown of tomosyn-1 inhibits exocytosis and

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Fig. 2A–D. The molecular machinery of insulin secretion. A Cellular events involved in glucose-stimulated insulin secretion. Reactions to the right (Electrical activity) are synonymous with the so-called triggering pathway in the beta-cell stimulus secretion coupling. Reactions to the left (Exocytosis) have the capacity to modify the beta-cell secretory response. Some of these reactions are likely to involved in the amplifying action of glucose on insulin secretion [124]. B SNARE proteins forming the beta-cell exocytotic core complex. C Mechanism for modulation of SNARE protein interactions by tomosyn-1. D The relative importance of electrical activity and exocytotic machinery in modulation of insulin secretion. Modulators are either enhancers (glucagon, Glg; glucagon-like peptide-1, GLP1; acetylcholine, ACh; pituitary adenylyl cyclase-activating peptide, PACAP; left) or inhibitors (galanin, Gal; somatostatin, Sst; adrenaline, Adr; right). The overall effects of the respective modulators are depicted by the large open bars, and the fractions that can be attributed to an action on electrical activity are shown by the smaller ﬁlled bars

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insulin secretion from INS-1E insulinoma cells, but does not affect transport and docking of insulin granules. Thus, tomoysin-1 appears necessary for fusion to occur [32] and/or priming of the granules for release [33]. As in other secretory cells, exocytosis in the beta cell is triggered by an elevation of the cytosolic free Ca2+ concentration. Capacitance measurements in conjunction with photorelease of caged Ca2+ have revealed a prominent component of exocytosis that proceeds at rates corresponding to 500 granules per second with a Kd as low as 17 mM [Ca2+]i. Such high Ca2+-concentrations are only attained in the immediate vicinity of the Ca2+ channels and probably involve a low-afﬁnity Ca2+-sensor. In addition, beta cells also exhibit a component of exocytosis that operates at Ca2+-concentrations as low as 0.17 mM and at rates as high as 3–4 granules per second [34,35]. The latter value is ∼109-fold higher than expected at this low [Ca2+]i for the low-afﬁnity pathway of release indicating that there are at least two Ca2+-sensing mechanisms controlling exocytosis in the beta cell. Exocytosis proceeding at low [Ca2+]i (“high afﬁnity component”) has been attributed to a small pool of insulin granules capable of release at low Ca2+-concentrations. It is accordingly referred to as the highly calcium-sensitive pool (HCSP). It seems likely that the Ca2+-sensor of beta-cell exocytosis, by analogy with other secretory cells, is a member of the synaptotagmin family. Fourteen mammalian isoforms have been described [36]. Recently, synaptotagmin IX has been proposed to represent the high-afﬁnity Ca2+-sensor in beta cells and silencing of this synaptotagmin by RNA interference inhibited glucose- and tolbutamide-induced insulin secretion in rat islets. It remains to be determined if synaptotagmin IX is also involved in exocytosis at high [Ca2+]i. Rapid exocytosis in chromafﬁn cells operate with a similar Ca2+-sensitivity as in beta cells (Kd = 13 mM). In these cells, synaptotagmin I functions as the lowafﬁnity Ca2+-sensor. However, this protein has been claimed not to be expressed in the beta cell [37]. Other putative Ca2+-sensors of exocytosis include piccolo. It has been shown to interact with the cAMP-sensing protein EPAC2, the sulfonylurea receptor SUR1 and the alpha1C-subunit of the Ltype Ca2+ channels [38], which all are essential for rapid Ca2+-induced exocytosis in the beta cell [14,39], and it is thereby well positioned to function as a low-afﬁnity Ca2+-sensor.

Biphasic Insulin Secretion Reflects Release of RRP Granules and Mobilization of New Granules from a Reserve Pool Ultrastructural data combined with biochemical determinations of insulin secretion and granule docking or the time course of capacitance increase in response to a step stimulus have led to the idea that the insulin granules exist

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in distinct functional pools and that these give rise to kinetically separable components of exocytosis [40,41]. A small fraction of the granules (1%) belong to a readily releasable pool (RRP) [2]. These granules are immediately available for release and give rise to the rapid component of capacitance increase, which lasts until the RRP has been depleted. Once the RRP is empty, the replenishment of RRP becomes rate-limiting for exocytosis. The RRP is reﬁlled by mobilization of granules from the reserve pool, which represents 99% of the granules in a beta cell. Granules belonging to this pool cannot undergo exocytosis and must undergo a series of preparatory reactions (“priming”) to attain release competence (Fig. 2A). As will be discussed below, this involves both chemical modiﬁcation of granules already in place at the release sites as well as physical translocation of granules towards the plasma membrane. Priming/mobilization is a time-dependent process and proceeds at a rate that is lower than the maximum rate of exocytosis. Accordingly, the capacitance increase in response to a step stimulus (i.e., photorelease of caged Ca2+) is biphasic and consists of rapid initial component followed by a sustained late component (Fig. 1D). It is tempting to propose that biphasic insulin secretion and exocytosis have the same cellular background. It should be noted, however, that the kinetics are very different: whereas the rapid component of capacitance increase is completed within <1s, 1st phase insulin secretion lasts ∼10 min. The peak rate of exocytosis (measured as granules released per second) indicated by the capacitance measurements is likewise much higher (>100-fold) than that observed in insulin release measurements. However, when allowance is made for the fact that exocytosis in intact islets (for reasons that remain unclear) is much slower than in isolated cells and the voltage dependence of secretion, a much better correlation is obtained [42]. Evidence supporting the hypothesis of a direct relationship between biphasic insulin secretion and pools of granules with variable release competence comes from the ﬁnding that both 1st phase insulin release and the rapid component of capacitance proceed well even in the absence of metabolic fuels, whereas 2nd phase release and the slower component of capacitance increase exhibit a strict metabolic dependence. There is also a reasonable quantitative agreement between the total number of granules belonging to the RRP and those undergoing release during 1st phase secretion. Further corroborating evidence was recently provided by the ﬁnding that genetic ablation of Munc13-1 selectively abolished 2nd phase insulin secretion and suppressed the slow phase of capacitance increase whilst leaving 1st phase release and the rapid component of capacitance release intact [43,44]. In neurons, loss of Munc13-1 results in complete suppression of synaptic transmission via emptying of the RRP of synaptic vesicles. These interesting data clearly illustrate the importance of granule priming for the development of 2nd phase insulin secretion. However, the fact that the initial component of

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capacitance increase was unaffected by loss of Munc13-1 demonstrates that this is not the sole mechanism for granule priming in the beta-cell and that alternative pathways, given enough time, are capable of ﬁlling the RRP to its full size. It is only during intense stimulation of secretion that the impairment becomes functionally signiﬁcant.

Molecular Machinery of Insulin Granule Trafficking Although the number of docked granules is sufﬁcient for several hours of glucose-stimulated insulin secretion, it is evident that exocytosis eventually must be balanced by the supply of new granules towards the plasma membrane. The ﬁrst glimpses of the cellular machinery involved in the intracellular granule trafﬁc were provided by the studies of Orci, Malaisse and their colleagues in the 1970s [45,46]. These studies, employing electron microscopy, documented the importance of the cytoskeleton for insulin granule transport and insulin secretion in general. However, electron micrographs can only provide a snapshot of the situation in the cell at the moment of ﬁxation. In the pioneering study of Kanazawa and colleagues [47], insulin granule trafﬁcking was monitored in living cells and in real-time using neonatal rat islet monolayers and phase contrast microscopy. Unfortunately this line of research was not pursued. The advent of novel cellular imaging techniques, such as confocal imaging and evanescent wave microscopy, combined with the development of a wide selection of ﬂuorescent proteins with enhanced spectral properties and the general advance in molecular biology, paved the way for more detailed cell physiological studies of the insulin granule trafﬁcking. These studies have revealed that insulin granule movements are of both random and directed nature [48]. The directed movements occur along the cytoskeleton (Fig. 3A). As in other spherical secretory cells, the beta-cell cytoskeleton is organized such that the mictotubule network occupies most of the cell with the exception of the space just beneath the plasma membrane which is occupied by the cortical actin network [48–50]. Confocal imaging of ﬂuorescent-labeled insulin granules in the insulinoma cell lines INS-1 and MIN-6 has revealed that insulin granule trafﬁc along the microtubule network is multidirectional and active even when the cell is not actively secreting. The velocity of moving granules exhibits extreme variability, reaching maximal velocities of 600–800 nm/s [48,51]. It is believed that the antegrade (forward) movement of granules depends on kinesin-1 [51]. A subset of very fast insulin granule translocations up to 1000 nm/s has been reported and attributed to kinesin-3, another member of the kinesin

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Fig. 3A–C. The molecular machinery of insulin granule transport. A An overall model demonstrating how microtubule-guided and actin-dependent insulin granule transport, as well as random granule movement (tortuous blue arrows) act in concert to replenish insulin granules at the plasma membrane. B Molecular motors kinesin-1 and dynein associated with microtubules in the beta cell. C Interactions between unconventional myosin 5a, synaptotagmin-like protein lacking C2 domains (Slac-2c), and the insulin granule (IG), as well as the actin network in the beta cell

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superfamily (Ivarsson and Renström, unpublished observations). Microtubule-guided movement in the reverse (retrograde) mode is, in beta cells as well as other tissues, driven by the dynein motor [52] (Fig. 3B). The actin ﬁlaments form a dense network beneath the plasma membrane. Disruption of this network accelerates Ca2+-induced exocytosis, indicating a “gate-keeping” function preventing hypersecretion of insulin. Granule passage through the actin web is regulated by the associated myosin motor proteins. Secretory cells typically express a set of different unconventional myosins. In the beta-cell myosin-5a [53,54] and myosin-6 are expressed but myosin-1c and non-muscle myosin-2 also seem to be present. Myosin-5a partially co-localizes with the insulin granules and knock-down of this protein suppresses regulated insulin exocytosis (Fig. 3C). Interestingly, myosin-5a appears to be particularly important for late phase secretion [54]. The cellular control of the directed insulin granule transport has not fully been elucidated. Signiﬁcant transport activity can be detected at substimulatory glucose concentrations but questions remain with regard to the Ca2+dependence and the identity of the trigger molecule(s). The directed movements have variably been reported to be Ca2+-dependent [55] and -independent [48], relying on Ca2+ entering from the exterior or mobilization from intracellular stores [56,57]. A regulatory role for AMPK controlling the kinesin-driven granule transport has also been reported [34]. A unifying model incorporating these disparate observations remains to be formulated. In addition to the directed movements, insulin granules also move by random diffusion within functional “cages” with a diameter of ∼0.9 mm. Diffusion itself is an unregulated process but it can be inﬂuenced by changes in viscosity in the surrounding medium that result, for example, from variations of the temperature. Our experiments indicate that diffusion of granules is essential for beta-cell granule trafﬁc. There is a strong correlation between the diffusion of granules within the cell and the frequency of directed granule translocations [48]. It is possible that the diffusional movements facilitate the interaction between the secretory granules and motorproteins at intracellular cytoskeletal junctions. In addition, reduced granule diffusion by cooling is associated with selective impairment of late phase insulin secretion [35,48,58].

Differential Control of Insulin Secretion by L- and R-Type Ca2+ Channels Inﬂux of extracellular Ca2+ through voltage-gated Ca2+ channels, with resultant elevation of [Ca2+]i, represents a key trigger of regulated insulin exocytosis. As discussed above, three subfamilies of voltage-gated Ca2+ channel

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exist: (1) L-type high voltage-activated (HVA) Ca2+ channel family that includes the CaV1.1, 1.2, 1.3, and 1.4 channels and that are sensitive to dihydropyridines (DHP [59–61]); (2) the non-L-type HVA channels CaV2.1 (P/Qtype), 2.2 (N-type) and 2.3 (R-type) that are blocked by omega-agatoxin IVA, omega-conotoxin GVIA, and SNX482, respectively [59,61,62]; and (3) the low voltage-activated (LVA) T-type Ca2+ channel family (CaV3.1, 3.2, and 3.3). The latter subtype differs electrophysiologically from the HVA Ca2+ channels in opening transiently upon modest depolarization [63,64]. In many cells, they fulﬁll a role as pacemaker [65]. The beta cell is equipped with a mixture of voltage-gated Ca2+ channels [15,16,66]. The presence of L-type Ca2+ channels in beta cells was established by radioisotopic and electrophysiological measurements more than 20 years ago [67] but the molecular identity of the beta-cell L-type Ca2+ channel has remained controversial. Whereas CaV1.1 and 1.4 are expressed exclusively in skeletal muscle and retina, respectively, CaV1. 2 and 1.3 are found in a wide variety of endocrine tissues. Knockout studies have shown that total CaV1.3 ablation leads to the suppression of insulin secretion; however, this effect is primarily due to a reduction in pancreatic beta-cell mass [68] and there is no evidence for a reduction of the L-type Ca2+ current in CaV1.3-deﬁcient mouse beta cells [69]. CaV1.2-null mice die at birth but beta-cell-speciﬁc knockout mice (beta-CaV1.2) exhibit an ∼50% reduction in the voltage-gated Ca2+ current in beta cells, similar to the inhibition of the beta-cell Ca2+-current obtained when the L-type Ca2+ channels are blocked pharmacologically [14]. Both electrical activity and glucose-induced [Ca2+]i-oscillations were essentially normal in CaV1.2-deﬁcient beta cells and islets. However, there was a strong (80%) inhibition of glucose-induced insulin release in isolated islets. Capacitance measurements revealed that rapid exocytosis (evoked by the ﬁrst two 500-ms depolarization to zero mV during a train) was almost abolished whereas exocytosis during the remainder of the train was much less affected. The results of these studies suggest that rapid exocytosis is tightly coupled to Ca2+-inﬂux via the L-type Ca2+ channels. There is evidence that the insulin granule and the L-type Ca2+ channel assemble into a functional complex via the synprint peptide [20,69]. It is therefore of interest that intracellular application of high concentrations of exogenous recombinant synprint peptide mimics the effects of knocking out CaV1.2 [69]. Dynamic measurements of insulin release using the perfused whole pancreas revealed that 1st phase insulin release was almost abolished in beta-Cav1.2-null mice,2 providing an

2 The control mice used in this study were heterozygous for the CaV1.2 knockout allele (+/−). However, no gene dose effect was observed and the magnitude of the Ca2+-currents was the same as in wild-type mice age.

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Fig. 4A,B. Granule movement and Ca2+ channel activity during phasic insulin secretion. A Stages in phasic insulin secretion: (i) prestimulatory mode, (ii) 1st phase insulin secretion, (iii) nadir phase, and (iv) 2nd phase. CaV1.2 Ca2+ channels are particularly important for early phase insulin release, whereas CaV2.3 channels primarily control 2nd phase by acceleration of mobilization. B Critical events in granule trafﬁc along microtubules (green) and the actin network (gray), as well as diffusional granule movement (tortuous blue arrows) in phasic insulin secretion (i–iv). In parallel, the dependence on Ca2+ inﬂux through L-type CaV1.2 (green) and R-type CaV2.3 channels during the different phases is also illustrated. The effects on granule trafﬁc are indicated by the width of the arrows (narrow arrow = slow; thick arrow = fast). (i) In the prestimulatory state, microtubuledependent granule trafﬁc is ongoing, but no exocytosis occurs at the plasma membrane. (ii) During 1st phase insulin granules close to L-type CaV1.2 channels are preferentially released. (iii) In the nadir phase, the pool of releasable granules at the plasma membrane is emptied, but Ca2+ inﬂux through R-type CaV2.3 channels activate mobilization, coinciding with the myosin-5a-dependent transport of insulin granules through the peripheral actin web and short-distance granule diffusion. (iv) During 2nd phase, microtubuledependent granule trafﬁc is at maximum; a new steady-state has been achieved favoring forward transport of granules to the plasma membrane, although both ante- and retrograde movements along microtubules are enhanced

additional argument for a correlation between biphasic capacitance increase and insulin secretion. It has been reported that the RIP-Cre construct commonly used to generate beta-cell-speciﬁc knockouts results in impaired glucose tolerance on its own [70]. It should be noted, however, that both the beta-Cav1.2-null and control mice used in the CaV1.2 studies described above contained only one RIPCre allele (i.e. RIPCre+/TG). We believe it is unlikely that the observed abnormalities result from non-speciﬁc effects of the RipCre transgene.

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Single-cell PCR measurements in isolated beta cells have conﬁrmed the expression of CaV1.2 in beta cells in addition to transcripts encoding P/Q, N and R-type Ca2+ channels [71]. Inhibition of R-type CaV2.3 channels by SNX482 (100 nM) or by a total knock-out reduce whole-cell Ca2+ currents by ∼25%, mirrored by a similar effect on insulin release in islets [72]. Pancreatic perfusion experiments showed that whereas 1st phase release was hardly affected, whereas 2nd phase insulin secretion was reduced. These distinct effects of L-type CaV1.2 and R-type CaV2.3 were also evident at the level of single-cell exocytosis. Taken together, these results suggest that the rapid 1st phase secretion reﬂects the release of insulin granules coupled to microdomains of elevated Ca2+ at the inner mouth of L-type CaV1.2 channels, whereas late 2nd phase secretion depends on R-type (CaV2.3 channels) (Fig. 4). However, it should be noted that the Ca2+-inﬂux via R-type Ca2+ channels does not appear to be coupled directly to exocytosis of a subset of beta-cell granules. If this were the case, then SNX482 and genetic ablation of CaV2.3 should inhibit insulin secretion/exocytosis to the same extent during the entire period of stimulation. This fact that these experimental procedures selectively inhibit 2nd phase insulin release instead favors the idea that Ca2+-inﬂux via R-type Ca2+ channels promotes granule mobilization. It has been demonstrated that a moderate global increase in [Ca2+]i, which in itself is insufﬁcient to cause insulin secretion, accelerates granule mobilization, and increases the size of RRP [73]. It seems possible that R-type Ca2+ channels mediate the Ca2+-inﬂux required for this component of release. Finally, the differential effects of SNX482 and the DHP-blockers nifedipine and isradipine on the kinetics of beta-cell exocytosis may indicate that R-type and L-type Ca2+ channels are spatially segregated and concentrate to different areas of the beta-cell plasma membrane.

How Do Granules Become Available for Release? Glucose inﬂuences insulin secretion by both triggering (i.e. involving closure of the KATP channels) and amplifying (i.e. post KATP channel closure) effects. Importantly, the latter pathway of release still requires induction of electrical activity and an elevation of cytoplasmic Ca2+. However, due to the amplifying action of glucose on secretion, the increase in Ca2+ produces a greater response. In a way analogous to the regulation of the KATP channel, the cytoplasmic concentrations of ATP and ADP have been found to reciprocally modulate exocytosis [74]. However, within the physiological range of concentrations, only ADP exerted an (inhibitory) effect. The latter effect was prompt and occurred within seconds after application of the nucleotide [75]. Changes in

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the ATP/ADP ratio at the plasma membrane, close to both the KATP channels and the release sites, may control both the triggering and amplifying actions of glucose. In addition to changes in the ATP/ADP-ratio, recent work has highlighted the role of the NADPH/NADP-ratio in the control of exocytosis. The effect was ATP-dependent, not mimicked by NADH/NAD and proposed to be mediated by the redox protein glutaredoxin and, possibly, thioredoxin [76]. A third hypothesis implicates glutamate as the intracellular messenger mediating the amplifying action of glucose on insulin secretion [77] but the role of this amino acid has been challenged [78]. Clearly, there is a long list of intracellular second messengers that could potentially mediate the amplifying action of glucose. It seems fair to conclude, however, that the jury is still out on the exact identity of the mediator although there are a number of strong candidates. It also remains unclear precisely how glucose via downstream metabolic products promotes exocytosis.

Proximal and Late Effects of Hormones and Neurotransmitters on Insulin Secretion Not just glucose but also many potentiators of insulin secretion act by both initiating electrical activity and stimulation of exocytosis. For example, GLP-1 (and other stimulators of adenylate cyclase like glucagon and forskolin) exerts a relatively minor effect on electrical activity (<20%) and yet exerts a very strong effect on insulin secretion (400%) [79,80]. Likewise, the strong stimulatory action of acetylcholine (ACh) on insulin secretion is only partly due to the enhancement of electrical activity [81,82]. For example, when applied in the presence of ∼10–15 mM glucose, ACh stimulates insulin secretion >5-fold. Only one-ﬁfth of this effect is accounted for by stimulation of electrical activity, the rest reﬂects stimulation of exocytosis. Although contribution by activation of protein kinase C, mediated by activation of phospholipase C and generation of diacylglycerol DAG with resultant enhancement of exocytosis [83] cannot be excluded, most of the latter effect appears to be secondary to the inositol trisphosphate (IP3-) evoked increase in [Ca2+]i [73]. This results in activation of Ca2+/calmodulin-dependent protein kinase 2, which in turn promotes granule priming and results in an increased RRP size. Conversely, many inhibitors of insulin secretion suppress insulin secretion more strongly than can be accounted for by the fairly mild effects on electrical activity. These are exempliﬁed by adrenalin, galanin, and somatostatin that, although they only transiently inhibit glucose-induced electrical activity in beta cells [23–25,84], almost abolish insulin secretion [84]. Like the effect on

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electrical activity, the latter effect involves a pertussis toxin-sensitive Giprotein. Using standard whole-cell recordings, the action of the inhibitory agonists on insulin secretion was found to be mediated by activation of the protein phosphatase calcineurin and intracellular application of speciﬁc inhibitors of this phosphatase prevented the inhibitory action of the agonists [26]. Interestingly, calcineurin is also involved in somatostatin-induced inhibition of glucagon secretion from pancreatic alpha-cells and in these cells catalyzes the depriming of secretory granules [85]. In chromafﬁn cells, calcineurin determines the size of the RRP via dephosphorylation of threonine residue 138 of the SNARE-protein SNAP25 [86]. Given that exocytosis of the insulin granules also involves SNARE proteins it seems possible that dephosphorylation of the same residue by calcineurin also explains the inhibition of insulin secretion by galanin and somatostatin in beta cells. These are just a few examples taken from our own work. However, the point that we wish to make is that modulation of exocytosis itself rather than more upstream processes, like electrical activity, represents a key mechanism controlling insulin secretion from beta cells (Fig. 2D). An interesting question is of course why the beta cell exhibits this dichotomy in the control of electrical activity and insulin granule exocytosis. Perhaps the answer lies in the existence of two parallel pathways of secretion in the beta cell involving LDCVs and SLMVs (see below). By controlling insulin release at the level of LDCV exocytosis, release of the SLMVs can proceed unchanged.

Two Pathways of Exocytosis Operate in Parallel in the Beta Cell Like neurons, many endocrine cells contain two classes of secretory vesicles [87]. Large dense-core vesicles (LDCVs) contain peptide hormones, whereas clear small synaptic-like microvesicles (SLMVs) store low molecular-weight neurotransmitters. SLMVs in beta cells are believed to accumulate gammaaminobutyric acid (GABA) [88], whereas the insulin-containing granules are believed to contain high concentrations of ATP. GABAA [89] and GABAB receptors [90] as well as purinergic receptors [91] have been identiﬁed in islets of Langerhans, suggesting that GABA and ATP, released by exocytosis of the SLMVs and LDCVs, can serve as paracrine/autocrine signaling molecules within the islet. Using beta cells engineered to express GABAA-receptor Cl− channels at high density as a biosensor (Fig. 5), depolarization- and Ca2+evoked GABA release has been reported [92]. This was taken as evidence of SLMVs capable of undergoing exocytosis, but recent ultrastructural data suggesting that the LDCVs may contain signiﬁcant amounts of GABA [93] makes

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Patch electrode

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Fig. 5. Receptor-based assay of beta-cell exocytosis. A beta cell is engineered to express ionotropic GABAA-receptor Cl− channels at high density. Stimulation of exocytosis of GABA-containing vesicles (SLMVs or LDCVs) (e.g., via intracellular dialysis with a high[Ca2+]i buffer) results in quantal release of gamma-aminobutyric acid (GABA). GABA thus secreted binds to the extracellular receptors and thus gives rise to transient membrane currents in the same cell. These can be recorded electrophysiologically using the same electrode as used for intracellular application of Ca2+. The transitory nature of the currents is due to diffusion of GABA away from the release sites and the receptors

it impossible to unequivocally conclude that SLMVs are capable of regulated exocytosis. Cell-attached capacitance measurements offer high resolution and permit the recording of capacitance increases as small as 10–20 aF (1 aF = 10−18 F). Using this technique, capacitance increases with an average step size as high 2.5 fF. In addition, a group of events with an average step amplitude as low as 0.15 fF has been observed [94] (Fig. 6A). Assuming spherical geometry and a speciﬁc membrane capacitance of ∼10 mF cm−2 these values predicts vesicle diameters of 0.3 mm and 80 nm, respectively. These are very close to the diameters of the LDCVs and SLMVs determined in ultrastructural analyses [95]. In many cases, vesicle fusion (seen as a capacitance increase) is transient and quickly followed by an endocytotic event of similar magnitude (Fig. 6B). Kiss-and-run exocytosis was quantitatively signiﬁcant and accounted for >25% of both the SLMV and LDCV exocytotic events. During kiss-and-run exocytosis, the secretory vesicle lumen is connected to the extracellular space via a lipid- and/or protein-lined fusion pore, through which the vesicular content exit [96]. We have estimated the dimensions of the fusion pore between the LDCV and SLMV lumens and the exterior by analysis of the patch conductance, which increases during exocytosis [97]. Fusion pores were detected during 60% and 80% of the SLMV and LDCV kiss-and-run events, respectively [98]. The average conductances were ∼50 pS for the SLMV and ∼200 pS for the

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Fig. 6. A Small (0.15 fF) and large (2 fF) capacitance steps presumably reﬂecting exocytosis of a synaptic-like microvesicle (SLMV) (i) and a large dense-core vesicle (LDCV) (ii). Im/2pf and Re reﬂect the capacitance increase (in aF or fF) and the associated conductance changes (in pS), respectively. Data from [94]. B Representative LDCV kiss-and-run event. The true vesicle capacitance (CV) and fusion pore conductance (Gp) were calculated from the Im and Re conductance components [98]. C Kiss-and-run vs “full” exocytosis as monitored by capacitance measurements. Exocytosis involves the opening of a fusion pore connecting the granule interior with the extracellular space. The conductance is detectable as long as the diameter (ø) of the fusion pore is <5 nm. When the diameter is about 1.5 nm (corresponding to a fusion pore conductance of ∼250 pS), insulin (black) remains trapped inside the granule whereas low-molecular-weight molecules like adenosine triphosphate or GABA (red) can escape. The electrophysiological deﬁnition of “full fusion,” when the fusion pore has dilated sufﬁciently to allow insulin exit, is based on fusion pore expansion beyond a detectable limit (∼10 nm diameter) and does not imply that the structure of the granule is lost. Fluorescence measurements have provided strong evidence that the integrity of the granule is maintained even after full fusion and release of the cargo [99,101]

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LDCV fusion pores. These conductances predict fusion pore diameters of 0.8 nm and 1.4 nm, respectively. Given that the insulin molecule has dimensions of 3.3 × 2.7 × 2.5 nm [99,100], it is clear that the LDCV fusion pore is too narrow to allow passage of monomeric insulin. Given these considerations it seems likely that during the majority of LDCV kiss-and-run events, insulin is retained within the vesicle. However, GABA, ATP and other low-molecularweight constituents of the secretory granules should be able to leave via the fusion pore. This would be consistent with work based on single-vesicle imaging of exocytosis in beta-cell utlilizing ﬂuorescently tagged granule membrane and cargo proteins, suggesting that the majority of exocytotic events are not associated with hormone release although the fusion pore is sufﬁciently open to allow pH equilibration [99–101]. However, there remain apparent discrepancies between the fusion pore properties established by the capacitance measurements and imaging. For example, a large body of data from ﬂuorescence imaging studies suggests that LDCVs remain structurally intact following exocytosis and that true “full fusion” of LDCVs is rare [99,101–106]. The fact that 80% of the exocytotic events detected by capacitance measurements were not kiss-and-run does not mean that the granule membrane collapses into the plasma membrane (but see [107] for a conﬂicting view). It only suggests that the fusion pore expansion beyond a detectable limit (∼10 nm diameter) (Fig. 6C). Despite these caveats, it is becoming increasingly clear that LDCVs do undergo kiss-and-run exocytosis. The role of LDCV kiss-andrun exocytosis is unlikely to be release of insulin. We favor the idea that these events provide a means of selective release of small molecules into the islet interstitium. For example, we have reported that ATP can be selectively released from LDCV without simultaneous release of the peptide cargo [101]. When combined with re-uptake mechanisms into the granules, it can be envisaged that the LDCVs can undergo several rounds of kiss-and-run exocytosis, thus providing a route for selective release of ATP into the islet interstitium where it may serve a paracrine function. The mechanisms of granule ATP reuptake remain to be determined. In chromafﬁn granules, ATP-dependent uptake of ATP into the granules has been documented. This uptake requires a positive intragranular membrane potential set up by a baﬁlomycin-sensitive proton pump and appears to involve DIDS-sensitive Cl− channels [108], but nothing is known about the situation in the beta-cell.

Type 2 Diabetes and Exocytosis Both insulin secretion and insulin action are impaired in type 2 diabetes (reviewed in [109]). Paradoxically, diabetic patients may have higher insulin levels than non-diabetic individuals but when allowance is made for the

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degree of hyperglycemia the insulin levels in diabetic patients are lower than in healthy controls. Both 1st and 2nd phase glucose-induced insulin release are reduced in type 2 diabetes [110]. We have argued elsewhere that a lowered cytoplasmic ATP/ADP-ratio due to an impaired glucose metabolism may contribute to this effect via suppression of granule priming [111]. In addition, studies on islets taken from the diabetic GK rat, a model of human type 2 diabetes, have revealed that impaired insulin secretory capacity correlates with reduced expression of key exocytotic proteins [112,113] and exhibits impaired exocytotic function [114,115]. Not just reduced expression of SNARE proteins but also mutations in these proteins may cause diabetes. For example, the blind drunk mutation (Bdr; (I67T)) of SNAP-25 is associated with impaired glucose tolerance in vivo and reduced glucose-induced insulin secretion in vitro [125]. Capacitance measurements revealed that the latter effect correlated with a reduction of exocytosis via interference with RRP replenishment. Micro-RNAs (miRNAs) are short (21–22 nucleotide) non-coding RNAs and are believed to be important regulators of transcription that act by promiscuous (i.e. more than one gene is affected) gene silencing. Currently, more than 300 miRNAs have been identiﬁed in the human genome. In silico analysis suggests that 20% of all genes may be regulated by miRNAs [116]. Overexpression of a pancreatic islet-speciﬁc microRNA (miR-375) in beta cells inhibits insulin exocytosis by mechanisms that are independent of alterations in the transmembrane Ca2+-ﬂuxes and intracellular Ca2+-signaling [117]. It is now important to determine if the expression levels of this miRNA as well as other islet miRNAs are altered in diabetic islets and how this in turn inﬂuences insulin secretion. As discussed above, imaging of exocytosis using ﬂuorescent markers has revealed that opening of the fusion pore (seen as pH equilibration and an increase in ﬂuorescence of pH-sensitive ﬂuorescent proteins associated with the interior of the granule membrane) is not obligatorily associated with release of the peptide cargo [101,104], presumably because the diameter of the fusion pore is too narrow to allow the exit of a bulky polypeptide. Processes that interfere with the expansion of the fusion pore can be envisaged to interfere with insulin secretion and cause diabetes. Indeed, as shown in a recent and potentially very important report [106], this appears to be the mechanism by which long-term culture at high glucose leads to the inhibition of insulin secretion. It was demonstrated that exposure of rat beta cells to 30 mM glucose for 48 h resulted in a dramatic increase in the fraction of kissand-run events (i.e., release events not associated with discharge of the peptide cargo) at the expense of the full fusions. If this occurs in man and at lower glucose concentrations, it is easy to envisage that a mild increase in

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plasma could glucose initiates a vicious cycle of progressive decrease in insulin secretion, elevation of blood glucose, etc. Type 2 diabetes also results in an elevation of plasma free fatty acid (FFA) levels [118]. Long-term exposure to lipids is associated with a marked suppression of glucose-induced insulin secretion and it has been hypothesized that this lipotoxic action contributes to the hyperglycemia/hypoinsulinemia of diabetes [119]. Lipids have been postulated to inhibit glucose-induced insulin secretion via uncoupling of mitochondrial metabolism [120,121]; the resultant decrease in ATP/ADP ratio impairs the ability of glucose metabolism to close the KATP channels which in turn causes loss of glucose-induced electrical activity and insulin secretion. However, our ﬁndings that long-term exposure of islets to FFAs, if anything, is associated with reduced KATP channel activity, stimulation of electrical activity, and no change in the ability of glucose to elevate [Ca2+]i, are not consistent with the hypothesis [122]. Indeed, insulin secretion stimulated by tolbutamide was reduced to the same extent as that evoked by glucose. This suggests that the lesion is at a level distal to KATP channel closure. However, both Ca2+ channel function and exocytosis were normal. The only parameter that was affected by FFAs was the insulin content which was reduced by >50%. Surprisingly, there was no corresponding decrease in the number of secretory granules, raising the interesting possibility that granular insulin content is not constant and may vary signiﬁcantly under different experimental conditions.

Concluding Remarks During the last 10–15 years there has been a remarkable advance in our understanding of the cellular and molecular control of insulin secretion. This is in part a consequence of the application of novel high-resolution techniques that allow insulin secretion to be monitored at the single-vesicle level and with millisecond temporal resolution. Our understanding of insulin secretion has also beneﬁted from the explosion of knowledge about the control of synaptic transmission and exocytosis in other endocrine cells (most notably the chromafﬁn cells). Modulation of insulin secretion involves both changes in beta-cell electrical activity and modulation of the exocytotic process itself; the latter modulation being perhaps particularly well developed in the beta cell. The data that have been generated on the regulation of exocytosis in the normal beta cell represents a valuable knowledge base that will facilitate the identiﬁcation of processes that malfunction under pathological conditions. A few possible examples have been highlighted here. Finally, it is important that we apply these high-resolution techniques to human beta cells. For example, it will be interesting to see how common gene polymor-

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phisms predisposing to diabetes (e.g., the one recently described in TCF7L2 [123]) correlates with exocytotic capacity. Acknowledgments. We thank Jenny Vikman and Rosita Ivarsson for help with Figs. 2 and 3, and Mrs. Nancy Lee for language editing. Supported by the Wellcome Trust, the European Foundation for the Study of Diabetes and the Swedish Research Council (SRC). E.R. is a Senior Researcher at the SRC; P.R. is a Wolfson-Royal Society Research Fellow.

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10. Mechanism of Insulin Exocytosis Analyzed by Imaging Techniques Shinya Nagamatsu and Mica Ohara-Imaizumi

Summary. Insulin is synthesized in rough endoplasmic reticulum (RER), delivered to the secretory granules where it is processed from proinsulin to insulin, stored, and released by the mechanism of exocytosis. Insulin release in vivo must be controlled minute by minute in order to maintain blood glucose levels within a narrow physiological range. The regulation of insulin exocytosis is critical to ﬁne control of glucose homeostasis. Insulin exocytosis consists of several steps, including docking, priming, and ﬁnal fusion with the plasma membrane. Fundamental components of secretory machinery such as soluble N-ethylmaleimide sensitive fusion protein attachment protein receptor (SNARE) play an important role in insulin exocytosis. Recently, several new techniques such as capacitance measurements, confocal, twophoton and total internal reﬂection ﬂuorescence (TIRF) microscopy have been applied to study insulin exocytosis. In particular, these new imaging techniques are powerful tools for addressing the mechanism of insulin exocytosis and examining the relationship between docking and fusion of granules and proteins implicated in the secretory machinery. This review focuses on the molecular mechanism of insulin exocytosis analyzed by the TIRF imaging system.

Introduction Beta cells of the endocrine pancreas play a critical role in the regulation of glucose homeostasis. These cells are dedicated to the biosynthesis and processing of proinsulin to mature insulin, which they then store in large dense core secretory vesicles (LDCV). Although beta cells contain a large number Department of Biochemistry, Kyorin University School of Medicine, 6-20-2 Shinkawa, Mitaka, Tokyo 181-8611, Japan

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of insulin secretory granules (about 13000 per cell) [1,2], only a fraction are released when the cell is stimulated. Glucose is the main signaling molecule that regulates not only insulin biosynthesis [3–6] and processing [7,8], but also insulin release [9,10]. The generally accepted model of glucose induced insulin release is as follows. Glucose metabolism generates a change in the adenosine triphosphate/adenosine diphosphate (ATP/ADP) ratio which initiates the downstream release of insulin. The change in ATP/ADP ratio causes ATP-sensitive potassium (KATP) channels to close, which prevents K+ ions from exiting the cell thereby leading to the depolarization of the plasma membrane [11,12]. Membrane depolarization leads to the opening of voltage dependent Ca2+ channels which initiates insulin secretion [13,14]. Insulin secretion stimulated by glucose displays a biphasic pattern [15,16] consisting of a rapidly initiated and transient ﬁrst phase followed by a sustained second phase. The biphasic release pattern is a consequence of the release of spatially and functionally distinct insulin granules [17] which are closely associated with proteins implicated in the secretory machinery. It is well known that fundamental components of the secretory machinery, such as soluble N-ethylmaleimide sensitive fusion protein attachment protein receptor (SNARE), which is required for the docking and fusion of vesicles in neuronal cells [18], are also expressed in pancreatic beta cells and play an important role in insulin exocytosis [19–22]. However, it is not clear how SNAREs regulate the biphasic pattern of insulin release. We have utilized total internal reﬂection ﬂuorescence (TIRF) imaging to examine the mechanism of biphasic release and the relationship between insulin secretory granules and SNAREs.

Visualization of Insulin Exocytosis by TIRF Microscopy The process of insulin secretion is shown in Fig. 1A. The secretory granules approach the plasma membrane from the intracellular space and dock to the plasma membrane with subsequent fusion, Thus, if the events taking place just beneath the plasma membrane are able to be imaged, valuable information concerning the mechanism of exocytosis may be obtained. Total internal reﬂection ﬂuorescence microscopy, which was originally developed by Axelrod [23,24], is a technique that meets this purpose. Total internal reﬂection ﬂuorescence microscopy provides a means to selectively excite ﬂuorophores in the cellular environment very near a solid phase (within <100 nm) without exciting ﬂuorescence from regions farther from the surface [23] (Fig. 1B). Fluorescence excitation by this thin zone of electromagnetic energy (called an “evanescent ﬁeld”) results in images with very low background ﬂuorescence virtually no out-of-focus ﬂuorescence, and minimal exposure of

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Fig. 1A,B. Insulin secretory granule exocytosis and total internal reﬂection ﬂuorescence (TIRF) microscopy. A Schematic drawing of the exocytotic process of an insulin granule. B Schematic drawing of the evanescent wave excitation. Total internal reﬂection occurs at incidence angles (q), which are greater than critical angle, thus producing an evanescent ﬁeld (highlighted area)

cells to light at any other planes in the sample. The TIRF imaging system allows small intensity changes to be interpreted as small motions of granules and dispersal of granule contents as exocytosis. The thin layer of illumination, “evanescent ﬁeld,” is produced by an excitation light beam traveling in a solid (e.g. a glass coverslip) incident at a high angle q upon the solid/liquid surface at which the cells adhere. The angle q must be large enough for the beam to totally internal reﬂect (TIR) rather than refract through the interface, a condition that occurs above some “critical angle.” Total internal reﬂection generates a very thin electromagnetic ﬁeld in the liquid with the same frequency as the incident light, exponentially decaying in intensity with distance form the surface. This ﬁeld is capable of exciting ﬂuorophores near the surface while avoiding excitation of a much larger number of ﬂuorophores. We

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applied this system to beta cells to visualize insulin secretion. The ﬁrst observation of the dynamic motion of insulin secretory granules was reported by Rutter’s group [25], where granules were labeled with a weak ﬂuorescent dye such as acridine orange which accumulates in acidic compartments with in cells. Acridine orange labels not only secretory granules but also other acidic compartments in cells. To speciﬁcally label insulin secretory granules, we generated an expression vector in which green ﬂuorescent protein (GFP) is fused to the C-terminus of preproinsulin (insulin–GFP). We later produced a recombinant adenovirus vector using the insulin–GFP construct. To acquire high-resolution real-time images of insulin granules, the TIRF microscope (Olympus Corporation, Tokyo, Japan) was equipped with a cooled chargecoupled device (CCD) camera (Andor Technology, Belfast, Northern Ireland), operated with Metamorph software (Universal Imaging, Downtown, PA). Using this system, we ﬁrst studied the dynamic motion of insulin granules in MIN6 insulinoma cells expressing insulin–GFP. The cells were stimulated with 50 mM KCl (Fig. 2) [26]. Insulin granule fusion was detected as a rapid and intense change of ﬂuorescence. Membrane depolarization by high KCl stimulation caused a prompt fusion of insulin granules docked at the plasma membrane. Recently, Chow’s group produced several kinds of probes to label the insulin granules and found that each fusion protein showed a different

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Fig. 2A,B. Total internal reﬂection ﬂuorescence image of green ﬂuorescent protein (GFP)labeled insulin granule motion during KCl (50 mM) stimulation. A Image observed before stimulation. Boxes indicate the granules docked to the plasma membrane, which will be fused with the plasma membrane by KCl stimulation. B Sequential images (1 × 1 µm, 300 ms intervals) of single granules. Adapted from Ohara-Imaizumi et al. [26]

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pattern of ﬂuorescence change during the fusion process [27]. In our system, the pattern of ﬂuorescence change of insulin granules is matched to that of endogenous insulin release measured by enzyme-linked immunosorbent assay. Thus, it appears that the behavior of our probe, insulin–GFP, is similar to that of endogenous insulin.

Biphasic Insulin Release Analyzed by the TIRF System Beta cells release insulin on glucose stimulation with a biphasic pattern that consists of a rapidly initiated and transient ﬁrst phase followed by a sustained second phase [28]. More than 30 years ago, Grodsky and colleagues proposed an insulin storage-limited mathematical model, the so-called two-compartment model, to explain the biphasic pattern [15,29,30]. Recently, Rorsman’s group performed elegant studies of insulin secretion using a combination of capacitance measurements and electron microscopy [2,14,16,31] and proposed a model for biphasic insulin release [17]. This model divides secretory vesicles into two distinct functional pools. One pool, the readily releasable pool (RRP) where granules gather near the plasma membrane, is releasecompetent and can undergo exocytosis. A second reserve pool of granules exists in the intracellular space to recruit granules to the RRP for replenishment. The RRP is estimated to contain 20–100 granules (0.2%–1.0% of total) and a substantial part of ﬁrst-phase insulin release could be attributable to exocytosis of RRP granules (1st phase). Once this pool of granules has been emptied, exocytosis proceeds at a lower rate, reﬂecting the low rate at which new granules are supplied for release by priming of reserve granules (2nd phase). Although this model provides a molecular explanation for Grodsky’s two-compartment model, it is based on indirect evidence, not direct observation of real-time insulin granule motion. Total internal reﬂection ﬂuorescence imaging has allowed us to directly observe the dynamic motion of single insulin granules undergoing exocytosis during biphasic insulin release in living beta cells. We transfected rat primary pancreatic beta cells with a recombinant adenovirus encoding insulin–GFP in order to label the insulin secretory granules and then examined the dynamic motion of granules near the plasma membrane [32]. Figure 3 shows a TIRF image of a single beta cell. The ﬂuorescence spots represent insulin granules docked to the plasma membrane (Fig. 3A). The beta cells were stimulated with high 22 mM glucose and sequential images of single granules were acquired every 50 ms. The data revealed the marked difference in exocytotic pathways between the 1st phase and 2nd phase. As shown in the sequential images (Fig. 3B) during the 1st phase (within the ﬁrst 4 min after the addition of 22 mM glucose), the fusing granules originated mostly from previously

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Fig. 3A–C. Total internal reﬂection ﬂuorescence images and analysis of single GFPlabeled insulin granule motion in rat primary beta cells. A TIRF image of GFP-labeled insulin granules close to the plasma membrane observed before glucose stimulation. B Sequential images of a granule docking and fusing with the plasma membrane during glucose stimulation (22 mM). During the 1st phase, the previously docked granules fuse with the plasma membrane, and during the 2nd phase, newcomer granules approach the plasma membrane and fuse. C Proposed model for biphasic insulin exocytosis mechanism. Adapted from Ohara-Imaizumi et al. [32]

docked granules that were visible before glucose stimulation. The ﬂuorescent spot suddenly brightened and vanished within 300 ms. On the other hand during the 2nd phase, the fusing granules arose from “newcomers”, which had been absent or only dimly visible before stimulation. It should be noted that the “newcomer” is found immediately after it reaches the plasma membrane. We plotted the number of fusion events versus time after stimulation. During the 1st phase, fusion occurs mostly from previously docked granules, whereas, in the 2nd phase, the fusing granules originate mostly from “newcomers.” Thus, direct imaging of granule motion in primary beta cells by our TIRF system show that Rorsman’s hypothesis [17] is basically correct, although some modiﬁcation of details of the 2nd phase may be required. Our

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data showed that during the 2nd phase, “newcomers” fuse directly without approaching the RRP. The depleted RRP is reﬁlled with new granules coming from inner pool that are not fused. Thus, it is likely that there is mechanistic difference between 1st and 2nd phase release. We propose the model for biphasic insulin secretion as shown in Fig. 3C. Recently Piston’s group reported that the reserve pool consists of two insulin granule populations and granule movement in both populations is involved in the cytoskeletal network [33]. Indeed, we found that translocation of granules from the reserve pool to the plasma membrane might be associated with the actin network (unpublished data). To visualize the translocation process of insulin granules in the cytosol, we are no now developing a new technology, so-called variable angle-TIRF (v-TIRF) microscopy [34]. A signiﬁcant advantage of v-TIRF microscopy is that it is able to visualize intracellular events more than 100 nm from the plasma membrane, which may enable us to acquire information along the z-axis (i.e., in three dimensions).

Site of Insulin Exocytosis Does insulin exocytosis occur anywhere in the plasma membrane or are there speciﬁc sites of insulin exocytosis? Recently, the basic components of the machinery that control the targeting and fusion of secretory vesicles with the plasma membrane were revealed (Fig. 4) [35,36]. Rothman and colleagues puriﬁed the proteins essential for vesicle transport and called these proteins SNAREs [37,38]. SNAREs are divided into two groups: (1) a vesicle-attached membrane protein (VAMP), so-called v-SNARE, and (2) two plasma membrane proteins, syntaxin and the 25-kDa synaptosomal-associated protein (SNAP-25), so-called t-SNAREs [38]. During the process of exocytosis, vSNARE pairs up with a unique cognate t-SNARE at the appropriate target membrane and this speciﬁc interaction docks the vesicles at the correct membrane site, with the subsequent membrane fusion. Because beta cells express v-SNARE, VAMP-2, and t-SNAREs, syntaxin 1A and SNAP-25 [19–22], insulin exocytosis is likely regulated by this protein–protein interaction (Fig. 4). We have recently developed a two-color TIRF system and to visualize the interaction between SNAREs, and insulin granules used it with the goal of determining the insulin granule exocytotic sites in living beta cells. For this purpose, we utilized a powerful protein transduction system [39,40]. The protein transduction domain (PTD) of the human immunodeﬁciency virus-1 (HIV-1) TAT protein has been shown to cross biological membranes efﬁciently and can be used to deliver peptides and proteins including antibodies (Ab) into cells. We produced a TAT-conjugated antibody against syntaxin 1A labeled with Cy3 [41]. Treatment of MIN6 insulinoma cells expressing insulin–GFP with

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Fig. 4. Soluble N-ethylmaleimide sensitive fusion protein attachment protein receptor (SNARE) proteins and SNARE complex formation. Upper panel Domain structure of SNARE proteins and organization of a SNARE-core complex. Lower panel v-SNAREs in the granule bind t-SNAREs in the plasma membrane, leading to the fusion process (modiﬁed from Chen and Scheller, 2001(18))

TAT-conjugated Cy3-labeled syntaxin 1 antibody enabled us to label endogenous syntaxin 1 in the plasma membrane of living cells. As shown in Fig. 5A, the Cy3-labeled syntaxin 1 clusters colocalize with the GFP-labeled insulin granule. We monitored the motion of GFP-tagged insulin secretory granules and their interaction with Cy3-labeled syntaxin 1 in live beta cells during stimulation. Stimulation with high KCl (50 mM) caused a marked membrane depolarization. The fusion events of insulin granules occurred exclusively at the syntaxin 1 clusters (Fig. 5A: granules to be fused are indicated by arrows). Figure 5B shows sequential images (1 × 1 mm, 300-ms intervals) of a single granule (green) and the syntaxin cluster (red) simultaneously observed during the stimulation. GFP-ﬂuorescent spots, which are present at the syntaxin clusters, suddenly brightened and ﬁnally spread as the GFP-tagged insulin diffused laterally on fusion of the secretory granule with the plasma membrane. No changes were observed in the syntaxin 1 clusters during and after granule exocytosis. Thus, the fusion of the insulin secretory granules appears to occur at the site of syntaxin 1 cluster at least during 1st-phase release.

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Fig. 5A,B. Total internal reﬂection ﬂuorescence image of GFP-tagged insulin granules and Cy3-labeled syntaxin clusters. A TIRF image of insulin granules (green) and syntaxin 1 clusters (red). Solid boxes represent the colocalization of insulin granules and syntaxin 1 clusters. B Sequential images of a single insulin granule (green) at the syntaxin 1 cluster (red) during 50 mM KCl stimulation. Adapted from Ohara-Imaizumi et al. [41]

It is well known that the release of neurotransmitters is restricted to a specialized presynaptic membrane compartment called the active zone, or hot spots [42]. Because the cytomatrix at the active zone (CAZ) has been implicated in deﬁning the site of Ca2+-dependent exocytosis of neurotransmitters [43], we hypothesized that a protein structurally related to the active zone associated protein, CAST, is a possible candidate for “hot spots” in beta cells [44]. CAST is not expressed in pancreatic beta cells [45]. However, CAST2, which is highly homologous to CAST, and known also as ELKS, is expressed in beta cells. We ﬁrst examined the localization of ELKS in beta cells [44]. As shown in Fig. 6, ELKS immunostaining was mainly observed in the pancreatic islets. Double staining for insulin and ELKS showed colocalization of ELKS and insulin. Confocal images showed that ELKS was localized at the plasmalemmal region of rat beta cells, especially those facing blood capillaries labeled with VE-cadherin, a marker for endothelial cells. These data suggest that ELKS clusters function as fusion sites for insulin secretory granules. We analyzed the interaction between the docking/fusion of GFPtagged insulin granules and ELKS clusters labeled by a TAT-conjugated, Cy3labeled ELKS antibody. Stimulation of beta cells with high KCl (50 mM)

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showed that fusion of insulin granules occurred at the ELKS clusters [44]. Thus, our data strongly suggest that ELKS plays a role in forming an active zone-like region in beta cells and fusion of insulin granules probably occurs on the site of blood capillaries.

Imaging Analysis in Diabetic Beta Cells Failure of insulin secretion, in particular 1st-phase insulin release, in response to glucose stimulation is a characteristic feature of type 2 diabetes [46,47]. Although there are several reports that have shown that abnormal glucose metabolism in diabetic beta cells contributes to the failure in insulin secretion [48–51], defects in the exocytotic machinery may also contribute to impaired insulin secretory response of diabetic beta cells. We examined the interaction of syntaxin-1 and SNAP-25 with insulin secretory granules from diabetic Goto–Kakizaki (GK) rats [52]. The TIRF image of insulin exocytosis in diabetic GK beta cells showed that fusion from previously docked granules on stimulation with 22 mM glucose was rarely observed during the 1st phase (Fig. 7). The number of fusion events during the ﬁrst phase is dramatically decreased in diabetic beta cells (Fig. 7B). Total internal reﬂection ﬂuorescence imaging of the immunostained insulin granules showed that there was a marked decline in the number of docked insulin granules (Fig. 7C,D). Because insulin content in GK beta cells is decreased [53], we calculated the number of granules docked to the plasma membrane adjusting for insulin content. The adjusted results still showed that the number of docked granules was decreased in diabetic GK rat beta cells to 57% of normal levels. Our results suggest that the failure in insulin release in diabetic beta cells could be due, at least in part, to impaired docking status. Because fusion of insulin granules occurs at the site of syntaxin 1A clusters during the 1st phase, we carried out the immunohistochemical analysis of syntaxin 1A and SNAP-25 localization in diabetic GK beta cells by TIRF microscopy. The TIRF images showed that the number of syntaxin 1 and SNAP-25 clusters in diabetic GK beta cells decreased to less than half those in normal beta cells (Fig. 8A,B). We also addressed the question of whether the normalization of blood glucose levels affects the number of t-SNARE clusters. If t-SNARE clusters determine the number of docked insulin granules, the change in number of t-SNARE clusters might correlate with that of docked granules. To test this idea, GK rats were treated with daily insulin injection for 2 weeks in order to normalize the blood glucose levels. We then examined the number of syntaxin 1 and SNAP25 clusters and that of docked insulin granules. As shown in Fig. 8, the number of t-SNARE clusters was increased as the number of docked insulin granules (Fig. 8C,D). Transfecting diabetic GK beta cells with recombinant

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Fig. 6A,B. CAST2 (ELKS) is present in pancreatic islet beta cells. A Colocalization of CAST2 and insulin in rat pancreatic islet. B Colocalization of CAST2, insulin, VE-cadherin, and syntaxin 1. Adapted from Ohara-Imaizumi et al. [44]

adenovirus encoding SNAP-25 increased the number of SNAP-25 clusters and docked insulin granules (Fig. 8A,B). Thus the number of t-SNARE clusters is correlated with that of docked insulin granules, which in turn are correlated with the number of fusion events from previously docked granules [52]. We speculate that, in diabetic GK beta cells, the decreased number of t-SNARE clusters results in impaired insulin granule docking status followed by a decreased fusion event that might lead to the loss of 1st phase insulin secretion. Genetic studies suggest that variation in the syntaxin 1A gene is associated with age at onset and insulin requirement in type 2 diabetic patients [54]. Furthermore, Ostenson et al. reported that expression of SNARE genes and proteins in pancreatic islets of type 2 diabetic patients is impaired [55]. Taken together, it is possible that decreased SNAREs (genetic or acquired)

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Fig. 7A–D. Total internal reﬂection ﬂuorescence images of insulin exocytosis in beta cells from diabetic GK rats. A TIRF image during 22 mM glucose stimulation, and sequential images of a granule docking and fusing during the 2nd phase. B The number of fusion events in 60-s intervals after glucose stimulation. C Epiﬂuorescence microscopy (EPIF) and TIRF imaging of docked insulin granules in diabetic beta cells. D The number of granules morphologically docked to the plasma membrane in diabetic beta cells was decreased compared to that in normal cells. Adapted from Ohara-Imaizumi et al. [32]

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Fig. 8A–D. Epiﬂuorescence microscopy and TIRF images of syntaxin, SNAP-25, and docked insulin granules in diabetic beta cells. Normal and diabetic beta cells were prepared and ﬁxed from GK rat pancreas, and immunostained with syntaxin 1, SNAP-25, and insulin antibodies. A EPIF and TIRF images of syntaxin and SNAP-25. B Histogram of the number of syntaxin 1 and SNAP-25 clusters. The number of syntaxin and SNAP-25 clusters in diabetic beta cells treated with insulin was restored to normal levels. C,D Images and the histogram of the number of insulin granules. Insulin treatment of GK rats and rescue of SNAP-25 in diabetic beta cells restored the number of docked insulin granules. Adapted from Ohara-Imaizumi et al. [52]

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expression affects the docking status of insulin granules, eventually leading to the impaired 1st-phase insulin secretory response.

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20. Nagamatsu S, Fujiwara T, Nakamichi Y, Watanabe T, Katahira H, Sawa H, Akagawa K (1996) Expression and functional role of syntaxin 1/HPC-1 in pancreatic beta cells. Syntaxin 1A, but not 1B, plays a negative role in regulatory insulin release pathway. J Biol Chem 271:1160–1165 21. Wheeler MB, Sheu L, Ghai M, Bouquillon A, Grondin G, Weller U, Beaudoin AR, Bennett MK, Trimble WS, Gaisano HY (1996) Characterization of SNARE protein expression in beta cell lines and pancreatic islets. Endocrinology 137:1340–1348 22. Nagamatsu S, Nakamichi Y, Yamamura C, Matsushima S, Watanabe T, Ozawa S, Furukawa H, Ishida H (1999) Decreased expression of t-SNARE, syntaxin 1, and SNAP-25 in pancreatic beta cells is involved in impaired insulin secretion from diabetic GK rat islets: restoration of decreased t-SNARE proteins improves impaired insulin secretion. Diabetes 48:2367–2373 23. Axelrod D (1981) Cell-substrate contacts illuminated by total internal reﬂection ﬂuorescence. J Cell Biol 89:141–145 24. Axelrod D (2001) Total internal reﬂection ﬂuorescence microscopy in cell biology. Trafﬁc 2:764–774 25. Tsuboi T, Zhao C, Terakawa S, Rutter GA (2000) Simultaneous evanescent wave imaging of insulin vesicle membrane and cargo during a single exocytotic event. Curr Biol 10:1307–1310 26. Ohara-Imaizumi M, Nakamichi Y, Tanaka T, Ishida H, Nagamatsu S (2002) Imaging exocytosis of single insulin secretory granules with evanescent wave microscopy: distinct behavior of granule motion in biphasic insulin release. J Biol Chem 277: 3805–3808 27. Michael DJ, Geng X, Cawley NX, Loh YP, Rhodes CJ, Drain P, Chow RH (2004) Fluorescent cargo proteins in pancreatic beta cells: design determines secretion kinetics at exocytosis. Biophys J 87:L03–L05 28. Nagamatsu S, Carroll RJ, Grodsky GM, Steiner DF (1990) Lack of islet amyloid polypeptide regulation of insulin biosynthesis or secretion in normal rat islets. Diabetes 39:871–874 29. O’Connor MD, Landahl H, Grodsky GM (1980) Comparison of storage- and signallimited models of pancreatic insulin secretion. Am J Physiol 238:R378–R389 30. Grodsky GM (1992) A threshold distribution hypothesis for packet storage of insulin and its mathematical modeling. J Clin Invest 51:2047–2059 31. Bokvist K, Eliasson L, Ammala C, Renstrom E, Rorsman P (1995) Co-localization of L-type Ca2+ channels and insulin-containing secretory granules and its signiﬁcance for the initiation of exocytosis in mouse pancreatic B-cells. EMBO J 14:50–57 32. Ohara-Imaizumi M, Nishiwaki C, Kikuta T, Nagai S, Nakamichi Y, Nagamatsu S (2004) TIRF imaging of docking and fusion of single insulin granule motion in primary rat pancreatic beta cells: different behaviour of granule motion between normal and GotoKakizaki diabetic rat beta cells. Biochem J 381:13–18 33. Hao M, Li X, Rizzo MA, Rocheleau JV, Dawant BM, Piston DW (2005) Regulation of two insulin granule populations within the reserve pool by distinct calcium sources. J Cell Sci 118:5873–5884 34. Wakazono Y, Sakurai T, Ohara-Imaizumi M, Nagamatsu S, Yamamoto S, Terakawa S (2006) Intracellular dynamics observed by mode switching of microscope with a light incidence to the interface at alternate angles through the ultra high NA objective. Proc SPIE 6088:449–454 35. Ferro-Novick S, Jahn R (1994) Vesicle fusion from yeast to man. Nature 370: 191–193 36. Rothman JE (1994) Mechanisms of intracellular protein transport. Nature 372: 55–63

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11. Two-Photon Excitation Imaging of Insulin Exocytosis Noriko Takahashi and Haruo Kasai

Summary. To elucidate the ﬁnal steps of secretion, we have established twophoton extracellular polar tracer (TEP) imaging, with which we can quantify all exocytic events in the plane of focus within intact pancreatic islets. We can also estimate the precise diameters of vesicles independently of the spatial resolution of the optical microscope and measure the fusion pore dynamics at nanometer resolution using TEP-imaging based quantiﬁcation (TEPIQ). During insulin exocytosis, it took 2 s for the fusion pore to dilate from 1.4 nm in diameter to 6 nm in diameter. Such unusual stability of the pore may be due to the crystallization of the intragranular contents. Opening of the pore was preceded by unrestricted lateral diffusion of lipids along the inner wall of the pore, supporting the idea that this structure was mainly composed of membrane lipids. TEP imaging has been also applied to other representative secretory glands, and has revealed hitherto unexpected diversity in spatial organization of exocytosis and endocytosis, which are relevant for physiology and pathology of secretory tissues. In the pancreatic islet, compound exocytosis was characteristically inhibited (<5%), partly due to the rarity of SNAP25 redistribution into the exocytosed vesicle membrane. Such mechanisms necessitate transport of insulin granules to the cell surface for fusion, possibly rendering exocytosis more sensitive to the metabolic state. TEP imaging and TEPIQ analysis will be powerful tools for elucidating molecular and cellular mechanisms of exocytosis and related disease, and to develop new therapeutic agencies as well as diagnostic tools.

Introduction Exocytic secretion is one of the fundamental cellular mechanisms for delivery of biosynthetic materials contained in cytosolic vesicles towards the extracellular space [1,2]. Exocytosis involves transport of secretory vesicles to target Division of Biophysics, Center for Disease Biology and Integrative Medicine, Faculty of Medicine, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan

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regions in the plasma membrane, docking of vesicles to the plasma membrane, and fusion of the two biological membranes, which may be followed by compound exocytosis on the fused vesicles and/or endocytosis. Regulation of exocytosis is handled by cytosolic Ca2−, cAMP, and GTPase in endocrine cells, exocrine cells, hematopoietic cells and neurons [3,4]. Key molecules involved in exocytosis have been identiﬁed during the last 15 years [5]. The fundamental mechanisms of these complex processes, however, are still largely unknown. Direct measurement of individual steps of exocytosis itself is still a matter of challenge. For instance, a key step of exocytosis, membrane fusion, is triggered by hemifusion of two biological membranes and opening of the nanometer fusion pore that connects extracellular space and vesicles. An ideal methodology should be able to track sizes and histories of vesicles, to reveal dynamics of fusion-pore formation and fates of omega-shaped fused vesicles, and to follow up compound exocytosis and endocytosis. Measurement of exocytosis in native and regenerating tissues is key to the development of new drugs and therapies. In this review, we describe twophoton excitation imaging, a powerful tool for studying exocytosis [6–14], and explain the importance of developing user-friendly two-photon microscope systems.

Two-Photon Excitation Imaging of Exocytic Events with Postfusion Labeling TEP Imaging A number of methods have been described for studying exocytosis and endocytosis (Fig. 1) including postfusion labeling, a method we developed that can be generally applied to any secretory preparation using a two-photon microscope (Fig. 1d). We gave this method the name “two-photon extracellular polar-tracer (TEP) imaging.” To visualize insulin exocytosis, we immersed the pancreatic islet preparations in a solution containing a polar ﬂuorescent tracer, sulforhodamine-B (SRB). The tracer was excited by laser-scanning microscopy with a mode-locked femtosecond-pulse Ti:sapphire laser (wavelength, 830 nm), revealing mesh structures inside the islets (Fig. 2a) [15]. The three-dimensional reconstructed images suggested that the ﬂuorescence reﬂected the microvasculature within the islets. When we stimulated the islets with a secretagogue such as 10–20 mM glucose, the exocytic event of individual insulin granules could be identiﬁed as the appearance of small ﬂuorescent spot (mean diameter: ∼0.4 mm full length at half maximum), reﬂecting the entry of extracellular polar ﬂuorescent tracer into the fused vesicle (Fig. 2b,c).

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Fig. 1. Analytical methods for studying exocytosis and endocytosis. TEP, two-photon extracellular polar tracer

Most of the ﬂuorescent spots disappeared within 10 s, reﬂecting full fusion of the vesicle membrane with the plasma membrane (Fig. 2c). We suggested that the transient ﬂuorescent spots represented individual exocytic events of insulin granules in the focal plane for the following reasons: (1) the diameter of the spots was similar to the diameter of insulin vesicles; (2) after chemical ﬁxation, the ﬂuorescent spots overlapped with the puncta that were positive by insulin immunohistochemistry; (3) the ﬂuorescent spots started to appear just as the cytosolic calcium concentration increased upon high glucose administration; (4) forskolin, which is well known to enhance insulin secretion, increased the frequency of appearance of the spots; and (5) the frequency of the appearance of spots could explain the amount of insulin secretion measured by radioimmunoassay [7,13].

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Fig. 2a–c. Two-photon extracellular polar tracer (TEP) imaging of insulin exocytic events in intact pancreatic islets. a Mouse pancreatic islet was immersed in a solution containing the polar ﬂuorescent tracer, sulforhodamine-B (SRB). TEP imaging revealed a mesh structure, which was tubular as shown in three-dimensional (3-D) reconstructed image. b,c Upon 20 mM glucose stimulation, an exocytic event of single insulin vesicle was detected as the appearance of a small ﬂuorescent spot (arrowhead in c)

Advantages of TEP Imaging TEP imaging can be readily performed using commercially available twophoton microscopes. The highly quantitative nature of TEP imaging takes advantage of a unique combination of the following three factors: (1) twophoton excitation imaging; (2) the use of extracellular polar tracers; and (3) the narrow intercellular spaces of intact tissues. Two-photon excitation is inherently resistant to light scattering in tissue preparations [16], allowing us to observe ﬁne structural alterations in the

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tissue without sacriﬁcing the spatial resolution of the microscope. In TEP imaging, we immerse the tissues in a solution containing a polar tracer, and one worries that the signal-to-noise ratio may be compromised because of the presence of large amounts of the tracers. This is not the case because the intercellular space of the tissues is normally as narrow as 20–40 nm, which is less than the diameter of most secretory vesicles (Fig. 1d). In addition, the intercellular space within a tissue is very clean, unlike the space between a cell and glass coverslip. Thus, the intercellular space provides one of the best experimental systems to study exocytosis, not to mention the fact that it is the physiological site for secretion. Two-photon excitation does not excite tracer molecules outside of the focal plane, and allows the use of high concentrations of polar tracers that are necessary to detect ﬁne alterations in cellular structures. With one-photon excitation, heat and phototoxic effects are generated in the entire tissue and damage the preparation. The photobleaching of tracers in the focal plane is effectively compensated for by diffusion of tracer molecules from outside of the focal plane. In conventional illumination, ﬂuorescent probes are continuously photobleached in the entire depth of preparations during image acquisition and bleaching cannot be overcome by diffusion. Therefore, absolute values of ﬂuorescence intensity cannot be utilized for quantization quantiﬁcation of exocytosis in conventional imaging. In one-photon excitation, the intensity of the actual excitation beam might be reduced by high concentrations of polar tracers present between the objective lens and the focal plane. This inner ﬁlter effect is particularly problematic when we examine structures deep with in the tissue, and when we perform multicolor imaging. In two-photon excitation, there is little absorption of the excitation beam except for the focal plane, and there is no inner ﬁlter effect for the excitation beam. We can control the state and concentration of the tracers in postfusion labeling, since the polar tracers are directly applied to the extracellular spaces. In contrast, it is difﬁcult to control the state of the probe in prefusion labeling (Fig 1c), since their properties are likely modiﬁed after they are incorporated into vesicles. Prefusion labeling is therefore not a good way to characterize exocytosis per se, although it is necessary to track the history of vesicle before exocytosis. Postfusion labeling can stain exocytic vesicles with simple diffusion without selection bias. The tracer SRB can ﬁll the vesicles, even small vesicles, fairly rapidly where the narrow fusion pore opens only transiently [12]. Thus, TEP imaging reveals all exocytic events in the visual ﬁeld [13], and can be used to study secretion without selection bias. Finally, two-photon excitation can simultaneously excite multiple tracers with single laser source as a consequence of the broader two-photon excitation spectra due to the blue shift in the two-photon excitation spectra relative

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to the simple prediction. The use of the same laser beam eliminates the offsets caused by chromatic aberration and misalignment of two laser beams. We do not use the pin hole in two-photon excitation but rather detect most of the ﬂuorescence with the photomultiplier. Thus, the collection of ﬂuorescent light is little affected by wavelength. Two-photon imaging enables completely simultaneous multicolor imaging without spatial offset. With the simultaneous multicolor imaging, we can measure precise time differences of the ﬂuorescent events, and perform ratiometric imaging to obtain internal states of molecule and cell. Exocytosis detected by TEP imaging is so deﬁnitive that the onset of fusion of individual vesicles can be deﬁned, the diameter of vesicle measured, and the fates of vesicles after exocytosis followed. TEP imaging can thus be used as stand-alone technique to study exocytosis and endocytosis. TEP imaging can be successfully combined with capacitance measurement [12] and electron microscopic investigation [11,12] when necessary.

Nanometer Measurement of Exocytic and Endocytic Vesicles TEPIQ Analysis Identiﬁcation of vesicle types is fundamental in studies of exocytosis since multiple types of exocytic and endocytic vesicles are functioning even within a single cell. The measurement of vesicle diameter is not a trivial problem because the diameters are close to or less than the spatial resolution of optical microscopes. TEP imaging allows us to measure diameters of vesicles by comparing ﬂuorescence intensity of individual exocytic events with that of the extracellular space (Fig. 3). TEP imaging can also characterize the fusion pore since exocytosis can be studied using multiple tracers with different colors. These approaches have been collectively termed TEP imaging-based quantitation (TEPIQ). Note that, if one-photon confocal imaging is used for TEPIQ-like analysis, the heat generated by the one-photon absorption throughout the tissue damages the tissues when we use sufﬁcient laser power to detect the small vesicles. TEPIQ analysis allows to be determined by three different methods. We can simultaneously perform them in the same preparation and thereby can conﬁrm the estimates.

TEPIQ Analysis of ∆V In TEP imaging, we acquire repetitive images only at one focal plane in order to follow the time course of exocytosis with is a rapid phenomenon. To

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Fig. 3. Three variations of TEP-imaging based quantiﬁcation (TEPIQ) analysis for estimating the diameter of exocytic vesicles

predict the volume of exocytic vesicles (∆V) from the stepwise increase in ﬂuorescence (FV) (Fig. 3), we therefore need to correct for z-axis resolution of a microscope as ∆V =

FV Fv = FE′ FE pxy ( 0)

where FE is the ﬂuorescence intensity per unit area in an xy-image of a solution with an inﬁnite depth, and F′E is the corrected ﬂuorescence of the extracellular solution [10]. The correction factor is called the efﬁciency of focal illumination, pxy(0), and can easily be obtained using small ﬂuorescent beads or quantum dot in each setup [10]. The volume thus obtained can easily be converted into vesicle diameter assuming sphericity of vesicles. In TEP imaging, the ∆V-TEPIQ analysis is always self-calibrating if images of exocytosis are acquired together with ﬂuorescence of an external solution. Full-width at half-maximal (FWHM) diameters of ﬂuorescence proﬁle of vesicles cannot be used to estimate diameters of vesicles less than 0.4 mm. Using TEPIQ analysis of ∆V, however, diameters of large dense-core vesicles of PC12 cells, beta cells and adrenal chromafﬁn cells have been estimated as 220 nm, 350 nm, and 500 nm, respectively, which are consistent with those reported by electron microscope.

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TEPIQ Analysis of ∆S We can also estimate the diameter of vesicles from the surface ﬂuorescence of vesicles stained with FM dyes, for example, FM1-43 (Fig. 3). The surface area (∆S) of vesicles is estimated from the stepwise increase in ﬂuorescence (FS) as ∆S =

FS FS = FM′ FM pxy ( 0)

where FM represents the membrane ﬂuorescence stained with the FM dye per square micron [10]. More precisely, FM is obtained from the total ﬂuorescence (FCell) of an xyz-image of an FM1-43-stained cell as FM = FCell/(4pR2) and R is the radius of the cell. This calibration experiment is tedious and even impossible when cells are not spherical. Instead, we measure the ﬂuorescence intensity per unit area (FC) of FM1-43 (20–25 µM) dissolved in the detergent CHAPS (40 mM), given that the emission spectrum of FM1-43 in CHAPS is similar to that of FM1-43 in the plasma membrane. We found that the value of FM was related to FC by the equation FM = mCFC, and the values of mC were similarly estimated as about 0.3 in beta cells and adrenal chromafﬁn cells [10,14]. Thus, the FM1-43 solution may be used to estimate the surface area. The value of FC should be determined daily to minimize the effects of changes in microscope settings. TEPIQ analysis of ∆S estimated the diameters of large densecore vesicles of PC12 cells, beta cells and adrenal chromafﬁn cells similarly as those of TEPIQ analysis of ∆V.

TEPIQ Analysis of ∆V/∆S TEPIQ analyses of ∆V and ∆S estimate the volume and surface area of an exocytic vesicle from the absolute ﬂuorescence intensity of an individual exocytic event. We can also obtain the diameter (fVS) of vesicles by taking the ratio between volume and surface areas, as

φvs = 6

∆V FV FM =6 ∆S FS FE

given that 4 6 × π r 3 / 4π r 2 = 2r = d . 3 TEPIQ analysis of ∆V/∆S estimated the diameters of large dense-core vesicles of PC12 cells and beta cells and adrenal chromafﬁn cells similarly as those of TEPIQ analysis of ∆V and ∆S [10,11,14]. Importantly, this multicolor correlational method can be applied even when individual events are not well resolved, and exocytic events are collectively measured. TEPIQ analysis of

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∆V/∆S have estimated the diameters of small vesicles, kiss-and-run endocytic vesicles, and clathrin-mediated endocytic vesicles of PC12 cells as 55 nm, 55 nm, and 90 nm, respectively [12]. These diameters were consistent with those estimated by electron microscopy using photoconversion of FM1-43 [12]. TEPIQ analysis of ∆V/∆S is insensitive to attenuation of the laser beam within the tissue. (If the same laser is used for excitation of both tracers any attenuation will result in the same reduction in the ﬂuorescence of both SRB and FM1-43.) TEPIQ analysis of ∆V/∆S also minimizes the out-of-focus effect [10]. Another advantage of TEPIQ analysis of ∆V/∆S is that it does not depend on the calibration constant, pxy(0). A disadvantage of TEPIQ analysis of ∆V/∆S is that it is highly dependent on the sphericity of the vesicle, becoming smaller when the vesicle is less spherical. Thus, TEPIQ analysis can be generally applied to measure exocytic and endocytic vesicles irrespective of a spatial resolution of the optical microscope.

TEPIQ Analysis of the Fusion Pore TEPIQ Analysis of Fusion Pore Diameters at Nanometer Resolution The fusion pore is the initial aqueous pore formed by fusion of two membranes, vesicle membrane and plasma membrane. Formation of the fusion pore is key to understanding the fusion reaction. Moreover, the fate of the fusion pore determines the fate of the vesicles. The fusion pore can dilate inﬁnitely to induce full fusion exocytosis, as we will describe below [7]. It can stably stay for a long time, and induce sequential exocytosis [11,14], or can be closed again for kiss-and-run or stay endocytosis [12]. Time-resolved membrane capacitance measurements have been used to estimate the initial diameter of the pore (Fig. 1a); however, they cannot estimate the diameters of pore larger than 2 nm and 6 nm for small and large vesicles [17], respectively, and may underestimate the pore size. Electrochemical (amperometric) measurements cannot by themselves estimate the diameter of the fusion pore (Fig. 1b). It was therefore useful to develop an imaging approach for measuring fusion pore diameter. We thus used ﬂuorescent polar tracers as nanometer-sized probes (Fig. 4a). If the pore is larger than the effective diameter of probe, the vesicle will be stained. We prepared solutions of two polar tracers, for example, SRB (0.3–0.7 mM) and dextrans of various molecular weights conjugated with ﬂuorescein (FD, 0.5–2 mM). Based on the molecular structures and light scattering, we estimate the hydrodynamic diameters of SRB and 10 kDa FD as 1.4 nm and 6 nm, respectively (Fig. 4a). We found that large dense-core vesicles of adrenal

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Fig. 4a–c. TEPIQ analysis of fusion-pore expansion in insulin vesicle. a Molecular dimensions of SRB and 10-kDa dextran conjugated with ﬂuorescein dextran (FD). The length of the second longest side of SRB (1.4 nm) is used for the effective diameter. b Time courses of single exocytic events of insulin vesicles stained with both SRB (red) and FD (green). c Schematic drawing of the expansion of the fusion pore

chromafﬁn cells and PC12 cells were nearly simultaneously stained with the two compounds within a time lag less than 50 ms, consistent with lifetime of the fusion pore of 10–50 ms in these cells studied with capacitance measurement and amperometry [18]. We also found that the fusion pore was reversibly closed without further opening to larger than 6 nm [7,11,14], which indicates that the pores are semistable and reversibly closed again. These results are consistent with the original notion of the fusion pore and prove that the fusion pore plays important role also in intact tissues. In beta cells, however, there was a signiﬁcant time lag between SRB and 10 kDa FD signals of 1–2 s with a mean value of 1.8 s (Fig. 4b,c) [7], suggesting that dilation of the fusion pore is exceptionally slow in insulin vesicles of beta cells. Consistent with this idea, the time course of staining of vesicles with polar tracers was slower than other cells (Fig. 4b). The following observations suggest that this was due to crystallization of insulin in the granules which prevented the dilation of the fusion pore. First, the pore dilation was further retarded by adding zinc (3 mM) to the extracellular solution [7], which is known to stabilize insulin crystal structure by binding to insulin [19]. Second, the pore dilation was signiﬁcantly faster in the islets of guinea pig, where crystallization of insulin is known to be weak [19]. Finally, ﬂattening of vesicles was found to start after the fusion pores were dilated to more than 12 nm, which was conﬁrmed by FD with molecular weight of 70 kDa. This ﬁnding

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may suggest that the ﬂattening of vesicles occurs when the fusion pore allows permeation of the insulin hexamer, which has a molecular weight of 36 kDa.

TEPIQ Analysis of Fusion-Pore Composition The slow dilation of the fusion pore of insulin vesicles allowed us to examine its molecular composition. It is worthwhile seeing the time course of staining of insulin vesicles with the lipidic dye FM1-43 since the pore size remained small for a few seconds. If the fusion pore is protein, the aqueous pore is the only pathway for FM1-43 staining (Fig. 5a, red). The staining of vesicles with FM1-43 should be similar or slower than SRB since FM1-43 accumulates in the plasma membrane. By contrast, if the fusion pore is lipid, FM1-43 can stain insulin vesicles via lateral diffusion from the already stained plasma membrane (Fig. 5a, green). Since the vesicle is as small as 350 nm, the lateral diffusion should be rapidly induced dependent on the geometry of the fusion pore. We found that FM1-43 stained the vesicle with an earlier onset and faster time course than SRB (Fig. 5b,c) [7,13]. More precisely, FM1-43 stained the

Fig. 5a–d. TEPIQ analysis of fusion-pore composition of insulin vesicle. a Two possible routes of staining of a vesicle with FM1-43. b Time course of single exocytic events of insulin vesicles stained with both SRB (red) and FM1-43 (green). c Schematic drawing of vesicle staining with FM1-43 and SRB. d Flux of FM1-43 molecule along the lateral wall of the fusion pore

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vesicle more than 64% when the SRB signal just started to appear. We estimated the diffusion constant of the FM1-43 molecule along the fusion pore (Fig. 5d) as 3.3 mm2/s [7]. This value is within the range for a pure lipid bilayer but is larger than that for the plasma membrane (<1 mm2/s), where lateral diffusion of membrane lipid is prevented by the actin-based membrane skeleton meshwork with the “picket” mechanism [20]. This indicates that the fusion pore is already lipid when the pore size is about 1.4 nm. This precludes the purely protein fusion pore model, and proteins, though present, do not disrupt the ﬂow of a lipophilic molecule, like FM1-43. The faster staining of vesicles by FM1-43 compared to SRB has been seen only in insulin granules, and not in the large dense-core vesicles of PC12 cells [11] or adrenal chromafﬁn cells [14]. This is because the pore expands rapidly in these cells and the vesicles are also rapidly stained with SRB.

Spatial Organizations of Exocytosis in the Pancreatic Islets Full-Fusion Exocytosis of Insulin Vesicles We conﬁrmed that TEP-imaging visualizes most insulin exocytosis in a focal plane of the islet, and the predicted amount of insulin secretion from the number of glucose-induced exocytic events can account for the amount of insulin release by radioimmunoassay [13]. Based on TEPIQ analysis, we strongly believe that the events reﬂect fusion of large dense-core vesicles of beta cells and can be used to characterize the properties of insulin exocytosis. Somewhat unexpectedly, exocytosis was observed all over the plasma membrane and there was only a slight inclination for exocytosis to occur preferentially towards the blood vessels [7]. This is consistent with the fact that no tight-junction structure is seen in islets and endocrine cells in general and indicates that the extracellular space is the major pathway for hormone secretion. We found that most fusion pores underwent full ﬂattening with the plasma membrane in 92% of glucose-induced events [7], though stable omega structures; the vesicles those were fused with the plasma membrane and whose fusion pores were open, were formed when a larger increase in cytosolic Ca2+ concentration was given. The full fusion exocytosis is the simplest form of exocytosis, and has been assumed for a long time, but TEP imaging has, for the ﬁrst time, deﬁnitively demonstrated its actual use in mammalian secretory cells. The full fusion occurred with two time constants, 1.5 s and 15 s in beta cells, somewhat slower than chromafﬁn cells, where its life was about 0.25 s [14]. As discussed below, this may be due to the time for dissolution of insulin crystal to hexamer. In fact, it has become evident that beta cells are rather exceptional in showing predominantly full fusion exocytosis, with two

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Fig. 6a–d. Two forms of compound exocytosis. a Sequential exocytosis is supported by lateral diffusion of t-SNAREs from the plasma membrane. b Multivesicular exocytosis may utilize two distinct t-SNAREs. c Full-fusion exocytosis, sequential exocytosis, and vacuolar sequential exocytosis in the pancreatic islet, acinus, and adrenal medulla, respectively. The green outline in the pancreatic acinus indicates actin coating. d Multivesicular exocytosis of eosinophils and basophils

other forms of exocytosis, sequential (2%) and kiss-and-run exocytosis (6%) being relatively infrequent. We believe that kiss-and-run exocytosis is suppressed in beta cells since such transient opening of fusion pore does little to insulin secretion. Protein kinase A further reduced kiss-and-run exocytosis in beta cells [13]. We think that sequential exocytosis (Fig. 6a) is also suppressed in beta cells in order for insulin exocytosis to be more sensitive to energy states of the cells [8]. One might also imagine that sequential exocytosis is infrequent in beta cells simply because most vesicles undergo full fusion and have no chance for secondary exocytosis. This is, however, not the case, since stimulation with caged-Ca2− compound greatly prolonged the lifetime of omega-shaped proﬁles, while it did not change the occurrence of sequential exocytosis [8].

Mode of Exocytosis Compound exocytosis is classiﬁed into two types. One type is sequential exocytosis, which we have observed in exocrine pancreas by TEP imaging [6]. The omega-shaped proﬁles in pancreatic acinar cells were stable for several minutes, and deep vesicles selectively fused with vesicles which had already fused with the plasma membrane (Fig. 6a,c). The chain reaction proceeded deep into the cytosol in an apparently random manner with the speeds of

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fusion not being slower in the deep cytosolic layer relative to the surface. We searched the literature, and found that we had observed a dynamic sequence of events whose static analysis was reported in 1965 with the electron microscope and named “sequential exocytosis” [21]. In endocrine tissues, sequential exocytosis was frequently observed in adrenal medulla (63% of the total exocytic events) [14] but is severely inhibited in pancreatic islet (2%) [8]. There is another type of compound exocytosis, called multivesicular (or multigranular) exocytosis, occurring when vesicles fuse to each other in the cytosol before fusing with the plasma membrane (Fig. 6b). This type of compound exocytosis is utilized by hematopoietic cells (Fig. 6d). Secretion can be explosive in multivesicular exocytosis (Fig. 6d) while secretion is under full control of agonist in sequential exocytosis (Fig. 6c). We think multivesicular compound exocytosis rarely happens in pancreatic islet or acinar cells, or in adrenal medulla, since we have observed the sudden appearance of compound vesicles (Fig. 6b). Note that the term “compound exocytosis” was used speciﬁcally for sequential exocytosis in an early study of mast cells [22], while “multivesicular exocytosis” was used in studies of eosinophils [23,24]. More recently, the term “compound exocytosis” has been used to include both sequential and multivesicular exocytosis [8,25]. Compound exocytosis was reported in studies of lactotrophs, alveolar type II cells, and mast cells without distinguishing sequential and multivesicular exocytoses. Both types of exocytosis exist in hematopoietic cells [23,24] and might be present in parotid acinar cells.

Lateral Diffusion of SNARE Proteins The fact that compound exocytosis was strictly sequential in pancreatic acinar cells [6], islet beta cells [8], adrenal chromafﬁn cells [14], and PC12 cells [11], even with a spatially homogeneous increase in cytosolic Ca2− concentrations, suggests that factors necessary for fusion diffuse from the plasma membrane into the vesicle membrane (Fig. 6a). For strict sequential exocytosis to happen, vesicles in the cytosol need to recognize the vesicles which are continuous with the plasma membrane for fusion, and the mechanism by which this recognition takes place is an important question in the ﬁeld of exocytosis. We have proposed that target SNARE proteins may diffuse from the plasma membrane to granule membrane and trigger sequential fusion events (Fig. 6a) [6]. This hypothesis was tested in islet beta cells [8] and in adrenal chromafﬁn cells [14]. Sequential exocytosis was prevalent and rapid in adrenal chromafﬁn cells, where lateral diffusion of SNAP-25 was detected in 43% of exocytic events

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and occurred with only with a short latency of 1.15 s from primary exocytosis. In contrast, lateral diffusion of SNAP-25 was detected in only 6% of exocytic events in the beta cells, where sequential exocytosis is slow and infrequent (2%) and occurred with a long latency of 6.5 s. Importantly, lateral diffusion of SNAP-25 was still detected in 54% of vesicles undergoing sequential exocytosis in beta cells, but only 5% in solitary exocytic events. Furthermore, when islets were treated with cyclodextrin to remove lipid rafts which might trap SNAP-25, sequential exocytosis was hastened and increased to 8.9% and lateral diffusion of SNAP-25 was detected in 15% of events [8]. Thus, lateral diffusion of SNAP-25 showed parallelism with the sequential progression of exocytosis in chromafﬁn cells and beta cells. Sequential exocytosis also involves other SNAREs, as redistribution of syntaxin-2 has been demonstrated by immunohistochemistry in pancreatic acinar cells. The assembly of the SNARE complex has often been considered to precede the events that trigger rapid exocytosis because SNARE assembly was reported to take more than 60 s in vitro [26]. Our data in chromafﬁn cells indicate that the complex formation could occur within in 1 s. Protein folding and assembling are known to be speeded up with molecular chaperons, and SNARE assembly may occur more rapidly in the cytosol than in vitro. It is important to identify the molecules that act as molecular chaperons for SNARE complex formation.

Perspectives We hope that the examples described in this review illustrate the power of TEP imaging in investigating exocytosis and endocytosis of secretory tissues and nervous systems. TEP imaging is the only method described to date that can be used to study secretion in intact tissues. It can be used to examine many important issues of exocytosis including vesicle diameter, fusion readiness, fusion-pore properties, life of omega-shaped proﬁles, swelling of vesicles, compound exocytosis, endocytosis, spatial organizations of exocytosis, as well as proteins involved in exocytosis, such as the SNARE proteins and the actin cytoskeleton. TEP imaging is also suited to studies of secretion in genetically modiﬁed mice, because we can study tissue preparations. For this reason, it can also be used to study secretion in human tissue samples. We can also utilize another powerful biophysical tool, namely caged compounds, in TEP imaging to quantify processes of exocytosis. It will be also possible in the future to perform simultaneous prefusion and postfusion labeling to track the entire life of vesicles before and after exocytosis. TEP imaging is emerging as a powerful method to study exocytosis just as patch-clamp methods are used to study ion channels. As TEP imaging becomes more widely used, the

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analysis will become automated. There is a bright future for TEP imaging in biomedical research.

References 1. Augustine GJ, Burns ME, DeBello WM, Pettit DL, Schweizer FE (1996) Exocytosis: Proteins and perturbations. Annu Rev Pharmacol Toxicol 36:659–701 2. Kasai H (1999) Comparative biology of exocytosis: Implications of kinetic diversity for secretory function. Trends Neurosci 22:88–93 3. Augustine GJ (2001) How does calcium trigger neurotransmitter release? Curr Opin Neurobiol 11:320–326 4. Seino S, Shibasaki T (2005) PKA-dependent and PKA-independent pathways for cAMP-regulated exocytosis. Physiol Rev 85:1303–1342 5. Jahn R, Lang T, Sudhof TC (2003) Membrane fusion. Cell 112:519–533 6. Nemoto T, Kimura R, Ito K, Tachikawa A, Miyashita Y, Iino M, Kasai H (2001) Sequential-replenishment mechanism of exocytosis in pancreatic acini. Nat Cell Biol 3: 253–258 7. Takahashi N, Kishimoto T, Nemoto T, Kadowaki T, Kasai H (2002) Fusion pore dynamics and insulin granule exocytosis in the pancreatic islet. Science 297: 1349–1352 8. Takahashi N, Hatakeyama H, Okado H, Miwa A, Kishimoto T, Kojima T, Abe T, Kasai H (2004) Sequential exocytosis of insulin granules is associated with redistribution of SNAP25. J Cell Biol 165:255–262 9. Oshima A, Kojima T, Dejima K, Hisa I, Kasai H, Nemoto T (2005) Two-photon microscopic analysis of acetylcholine-induced mucus secretion in guinea pig nasal glands. Cell Calcium 37:349–357 10. Kasai H, Hatakeyama H, Kishimoto T, Liu TT, Nemoto T, Takahashi N (2005) A new quantitative (two-photon extracellular polar-tracer imaging-based quantiﬁcation (TEPIQ)) analysis for diameters of exocytic vesicles and its application to mouse pancreatic islets. J Physiol 568:891–903 11. Kishimoto T, Liu TT, Hatakeyama H, Nemoto T, Takahashi N, Kasai H (2005) Sequential compound exocytosis of large dense-core vesicles in PC12 cells studied with TEPIQ (two-photon extracellular polar-tracer imaging-based quantiﬁcation) analysis. J Physiol 568:905–915 12. Liu TT, Kishimoto T, Hatakeyama H, Nemoto T, Takahashi N, Kasai H (2005) Exocytosis and endocytosis of small vesicles in PC12 cells studied with TEPIQ (two-photon extracellular polar-tracer imaging-based quantiﬁcation) analysis. J Physiol 568: 917–929 13. Hatakeyama H, Kishimoto T, Nemoto T, Kasai H, Takahashi N (2006) Rapid glucose sensing by protein kinase A for insulin exocytosis in mouse pancreatic islets. J Physiol 570:271–282 14. Kishimoto T, Kimura R, Liu TT, Nemoto T, Takahashi N, Kasai H (2006) Vacuolar sequential exocytosis of large dense-core vesicles in adrenal medulla. EMBO J 25: 673–682 15. Takahashi N, Nemoto T, Kimura R, Tachikawa A, Miwa A, Okado H, Miyashita Y, Iino M, Kadowaki T, Kasai H (2002) Two-photon excitation imaging of pancreatic islets with various ﬂuorescent probes. Diabetes 51 Suppl 1:S25–S28 16. Dong CY, Koenig K, So P (2003) Characterizing point spread functions of two-photon ﬂuorescence microscopy in turbid medium. J Biomed Opt 8:450–459

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17. Klyachko VA, Jackson MB (2002) Capacitance steps and fusion pores of small and large-dense-core vesicles in nerve terminals. Nature 418:89–92 18. Breckenridge LJ, Almers W (1987) Currents through the fusion pore that forms during exocytosis of a secretory vesicle. Nature 328:814–817 19. Dodson G, Steiner D (1998) The role of assembly in insulin’s biosynthesis. Curr Opin Struct Biol 8:189–194 20. Fujiwara T, Ritchie K, Murakoshi H, Jacobson K, Kusumi A (2002) Phospholipids undergo hop diffusion in compartmentalized cell membrane. J Cell Biol 157:1071– 1082 21. Ichikawa A (1965) Fine structural changes in response to hormonal stimulation of the perfused canine pancreas. J Cell Biol 24:369–385 22. Alvarez de Toledo G, Fernandez JM (1990) Compound versus multigranular exocytosis in peritoneal mast cells. J Gen Physiol 95:397–409 23. Scepek S, Lindau M (1993) Focal exocytosis by eosinophils—compound exocytosis and cumulative fusion. EMBO J 12:1811–1817 24. Hafez I, Stolpe A, Lindau M (2003) Compound exocytosis and cumulative fusion in eosinophils. J Biol Chem 278:44921–44928 25. Pickett JA, Edwardson JM (2006) Compound exocytosis: mechanisms and functional signiﬁcance. Trafﬁc 7:109–116 26. Fasshauer D, Antonin W, Subramaniam V, Jahn R (2002) SNARE assembly and disassembly exhibit a pronounced hysteresis. Nat Struct Biol 9:144–151

III. Pancreatic Development and Beta-Cell Formation

12. Regulation of Beta-Cell Growth and Death Christopher J. Rhodes

Summary. Pancreatic beta-cell mass, under normal circumstances, is maintained at an optimal level to provide for a normal metabolic load. It should not be viewed as static, and slow in turning over. Indeed, the adult beta-cells adults are plastic and able to increase their population to adapt to changes in metabolic demand, such as in pregnancy or nondiabetic obesity. Net changes in beta-cell mass are reﬂective of the amount of growth (i.e., the sum of betacell replication, neogenesis and size) minus the degree of beta-cell death (i.e., the sum of beta-cell apoptosis, necrosis and autophagic cell death). In some circumstances, such as prolonged obesity and insulin resistance, the beta-cell works under pressure trying to meet the metabolic demand, but eventually succumbs to a collective number of stresses that lead to an increase in betacell death, a subsequent reduction in beta-cell mass, and the eventual onset of type 2 diabetes. In this chapter, what is currently known about mechanisms of beta-cell growth and death are looked at in detail, with an emphasis on adaptive mechanisms in obesity and pregnancy and the signal transduction pathways involved in the control of beta-cell growth. Also, the various stresses on the beta cell that may lead to the onset of type 2 diabetes will be outlined and the inﬂammatory signaling pathways that contribute to increased betacell apoptosis/necrosis examined.

Introduction Insulin, produced in islet beta cells of the endocrine pancreas, is the most important mammalian anabolic hormone for the control of normal metabolic homeostasis. Under normal circumstances the population of beta cells is maintained at an optimal number to meet the metabolic demand. Yet, Department of Medicine, Section of Endocrinology, Diabetes, and Metabolism, University of Chicago, 5841 S. Maryland Avenue, MC 1027, Room N138, Chicago, IL 60637, USA

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beta-cell mass should not be considered static, as it exhibits plasticity to meet natural changes in metabolic load, as in pregnancy or obesity [1]. Failure of the beta-cell mass to adapt to changes in metabolic demand can be devastating, leading to an insulin deﬁcient state and the onset of type 2 diabetes [1,2]. The concept of insufﬁcient beta-cell mass as the key factor in the pathogenesis of type 2 diabetes has only recently been widely acknowledged. In this regard, it is worth noting a conceptual similarity between type 2 and type 1 diabetes, the latter also being characterized by reduced beta-cell mass due to an autoimmune destruction of beta cells [3]. As such, diabetes mellitus should be primarily considered a pancreatic beta-cell disease. In this chapter, the focus will be on the regulation of beta-cell growth and mechanisms that initiate beta-cell death related to the pathogenesis of type 2 diabetes. The autoimmune pathogenesis of type 1 diabetes will not be considered here in detail, but interested readers should refer to excellent reviews on the subject [3,4]. In addition, it is acknowledged that insulin secretory dysfunction [5–7], along with loss of beta-cell mass, also contributes to the pathogenesis of type 2 diabetes.

Pancreatic Beta-Cell Growth General Pattern of Normal Postnatal Beta-Cell Growth Most of the information on postnatal beta-cell growth has come from studies in rodents. There is a small proviso in using animal models relative to humans. However, the data that has emerged to date from studies in humans (albeit small) [8–13], suggest similar processes occur in rodents and humans, at least qualitatively. A normal neonate has a certain population of differentiated pancreatic islet beta cells that have developed in utero. But this is limited and insufﬁcient for life outside the womb. Quite soon after birth there is a rapid rise in beta-cell mass due to a marked increase in both beta-cell replication and neogenesis [1,14,15] (Fig. 1A). However, this early burst of beta cell growth is transient and begins to tail off soon after weaning [14]. There is also a slight increase in beta cell apoptosis at this time, but this is far outweighed by the replication and neogenesis so that there is a net a net increase in beta-cell growth [14]. This apoptosis is probably symptomatic of some cellular rearrangement going on during this time, associated with the islet cell neogenesis. Thereafter, in the normal situation, the beta-cell mass stays relatively constant with very low rates of beta-cell (estimated at ≤0.2%) replication or apoptosis, and negligible neogenesis [14,16]. Interestingly, there does not appear to be much of an increase in beta-cell mass (or beta-cell replication and neogenesis) during puberty (Fig. 1A). This implies that a normal individual may have a relative excess of beta

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Fig. 1. Hypothetical changes in beta-cell growth during a human lifespan. Net changes in pancreatic beta-cell mass are a reﬂection of the amount of growth (i.e., the sum of beta-cell replication, neogenesis, and size) minus the degree of beta-cell death (i.e., the sum of beta-cell apoptosis, necrosis, and autophagic cell death). This ﬁgure represents current thinking of changes in beta-cell mass (upper panels) in a human lifespan derived from limited human data and that from rodents. The changes in beta-cell mass are subdivided into changes in beta-cell growth (middle panels) and beta-cell death (lower panels). A depicts changes in beta-cell mass under normal circumstances (solid line); the increase in beta-cell mass in nondiabetic obesity (dotted line), and the decrease in beta-cell mass found in obesity-linked type 2 diabetes (dot/dash line). Details can be found in the text

cells early on in life and grows into their beta-cell mass as they proceed into adulthood. Setting a baseline beta-cell population early on in childhood establishes a critical mass from which to expand in response to increased metabolic load, such as in pregnancy or obesity. This concept might have implications for a susceptibility to the onset of type 2 diabetes (see below).

General Mechanisms of Beta-Cell Growth Postnatal beta-cell growth is complex and contributed by several mechanisms, including replication, neogenesis, size and transdifferentiation. These are considered individually below.

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Fig. 1. B depicts changes in beta-cell mass for a mother who has two children. Two bursts of increased beta-cell mass in adulthood are observed to compensate for each of the two developing fetuses (upper panel). This is linked to increases in beta-cell growth (middle panel) and, after birth of the child, followed by increases in beta-cell apoptosis that bring the beta-cell mass back to normal levels (lower panel)

Beta-Cell Replication Beta-cell replication is the primary means of increasing the beta-cell population. Several peptide growth factors and nutrients, including glucose, have been shown to increase the rate of beta-cell replication [2], but these observations are based mostly on in vitro studies and it is essentially unknown what drives change in beta-cell mitogenesis in vivo. However, it is known that regulatory components of the cell-cycle are essential for normal beta-cell replication, especially cyclin-D2 [17,18] and cyclin-D dependent kinase-4 (CDK-4) [19]. Cyclin-D2 knockout mice have markedly smaller islets and consequently deﬁcient in beta-cell mass causing glucose intolerance at only 2 weeks of age [17,18]. Likewise, CDK-4 knockout mice appear normal at birth but have marked difﬁculties increasing beta-cell mass and consequently have insulin dependent diabetes [19]. In contrast, transgenic expression

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of a deﬁciently regulated form of CDK-4 causes beta-cell hyperplasia and hypoglycemia [19]. These studies indicate the great importance of the neonatal burst in beta-cell replication in establishing an adequate beta-cell mass. A more detailed outline of the molecular mechanisms involved in beta-cell replication can be found in the chapter by Salpeter and Dor. Many factors and nutrients can induce beta-cell replication based mostly on in vitro studies [2]. The most prominent factors shown to increase beta-cell replication are glucose, insulin-like growth factor-1 (IGF-1), growth hormone, hepatocyte growth factor (HGF), prolactin, and placental lactogen. But it is unclear which are the physiological factors that drive speciﬁc beta-cell replication in vivo, especially during the early neonatal burst of beta-cell replication that is the major contributor to deﬁning beta-cell mass in adulthood. Beta-Cell Neogenesis Beta-cell neogenesis describes the formation of new beta cells from the ductal epithelium of the pancreas [14,20,21]. It has been suggested that beta-cell neogenesis might not occur in mice [22], yet this has been documented in rats and humans [9,23]. However, it should be noted that, for the most part, beta-cell neogenesis is detected by “insulin-positive” staining cells associated in pancreatic ducts in pancreatic sections, and caution should be taken as this does not necessarily mean that these are “insulin-producing” cells [24,25]. Moreover, care should be taken with this sort of analysis since a single “insulin-positive” cell might actually be the tip of an iceberg and a peripheral part of an actual islet that is revealed when one looks further into the pancreatic section [25]. This emphasizes the importance of conducting a serial section analysis of ﬁxed pancreata when investigating beta-cell neogenesis. Although it is generally accepted that beta-cell neogenesis likely occurs, the mechanism is poorly deﬁned. It is probable that the process of beta-cell neogenesis will be complex with mature beta cells developing from adult pancreatic stem cells through a series of intermediate precursor cells, conceptually similar to what occurs for development of beta cells embryologically. However, it is likely that adult beta-cell neogenesis and embryological beta-cell development have quite different mechanisms and factors driving them at each cellular differentiation step. Unveiling the process of beta-cell neogenesis will be difﬁcult. Currently, no bone ﬁde cellular markers of pancreatic stem cells and/or intermediate precursor cells are known for beta-cell neogenesis. Carbonic anhydrase-II has recently been proposed as a ductal precursor cell marker [26], and shows promise but has yet to be conﬁrmed. However, care must be taken in deﬁning beta-cell precursor cell markers, since most of those previously proposed, such as nestin [27], have subsequently been shown not to follow a beta-cell lineage [28,29]. Indeed, much needed lineage tracing studies are

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currently being conducted to better deﬁne the process of beta-cell neogenesis and subsequently these should reveal the identity of speciﬁc markers at each step in the beta-cell differentiation process. These markers should then be useful in better deﬁning the process of beta-cell neogenesis in humans, where lineage tracing studies are obviously impossible. As such, in the next coming years beta-cell neogenesis mechanisms will be much better deﬁned. As with the process of beta-cell neogenesis itself, factors that trigger this process are also not well deﬁned. However, IGF-1 [30], the incretins glucagon-like peptide-1 (GLP-1) and glucose-dependent insulinotropic polypeptide (GIP) [31–34], and the combination of gastrin and epidermal growth factor (EGF) [35] have also been implicated in increasing beta-cell mass in vivo, in part by increasing beta-cell neogenesis. Beta-Cell Size The volume of a beta cell also contributes to the size of the beta-cell mass. However, it is unknown whether a bigger beta cell actually means a larger capacity to produce, process, store, and/or secrete insulin, although this is often assumed to be the case. Physiological studies and electron microscopy (EM) analysis of big versus small beta cells is needed to better establish this notion. It is also unclear how beta-cell size is controlled, although this is likely partly linked to the regulation of general protein synthesis via the serine/ threonine protein kinases mTOR and p70 ribosomal S6 protein kinase (p70S6K) [36,37]. In this regard, p70S6K knockout mice have a slightly reduced beta-cell mass due decreased beta-cell size, and consequently have a reduced insulin secretory capacity and mild glucose intolerance [38]. Beta-Cell Transdifferentiation Beta-cell transdifferentiation is a term used to imply a transformation of a pancreatic exocrine acinar cell into pancreatic endocrine beta cell [25]. Although there are some in vitro studies to support this notion [39–42], and some do not [43], convincing in vivo evidence is lacking. Moreover, a proviso should be made in that many of the in vitro studies have been conducted in polyclonal transformed exocrine cell lines (Panc-1 and AR42J-cells) that also contain a minority of insulin positive beta cells within them, so it is difﬁcult to access beta-cell replication versus transdifferentiation. Indeed, the many difﬁculties associated with studying beta-cell neogenesis described above also apply to studies of beta-cell transdifferentiation. This includes the speculation that a single beta-cell present in a sea of exocrine cells on a pancreatic section has arisen via transdifferentiation [25]. Again, this could simply be a beta cell on the periphery of an islet that would be revealed by a more thorough serial section analysis. Lineage tracing studies in transgenic animal models are much needed to settle the controversy of whether beta-cell trans-

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differentiation occurs in vivo or not [44]. One recent study suggests the latter [45]. Indeed, this has been conﬁrmed by a second study that clearly implicates pancreatic exocrine cells do not “transdifferentiate” into pancreatic endocrine beta cells in vivo [46]. It should also be noted that several groups have shown that liver derived cells can be transdifferentiated into “insulin expressing cells” by various genetic manipulations [47,48]. However, while this might hold some future promise of using gene transfer technologies to generate beta cells therapeutically, it does not occur in the normal physiological setting.

Physiological Circumstances for Increasing Beta-Cell Growth Pancreatic beta-cell mass, under normal conditions in adulthood, remains relatively constant and is optimal to cope with the metabolic demand (Fig. 1A). However, certain physiologically relevant circumstances raise the metabolic load and the beta-cell mass correspondingly increases to compensate. There are two major situations where this occurs, obesity (Fig. 1A) and pregnancy (Fig. 1B). Pregnancy The developing fetus during pregnancy places an increased metabolic demand on the mother. As a consequence, the mother’s pancreatic beta-cell mass approximately doubles [49] (Fig. 1B). It is known that this enhancement of beta-cell mass is mediated by an increase in replication and cell size [49], but it is unclear whether this is also contributed by a rise in beta-cell neogenesis. Out of all the growth factors proposed to increase beta-cell mass [2], those that drive it in pregnancy are best documented. They are prolactin and placental lactogen [49]. Intriguingly, after birth the beta-cell mass rapidly decreases back to normal via a regulated increase in beta-cell apoptosis [50] and reduction in beta-cell size (Fig. 1B). It is unknown how this post-partum burst of beta-cell apoptosis is controlled. Nondiabetic Obesity The increase in adiposity, as well as other factors affecting general homeostasis in obesity, including insulin resistance, also increase the metabolic load and consequently the demand for insulin. However, this is usually compensated for by an increase in beta-cell mass [10,51] (Fig. 1A). As such, at least two-thirds of obese human subjects do not enter a state of insulin insufﬁciency to develop obesity-linked type 2 diabetes. This increase in beta-cell mass found in nondiabetic obesity is contributed to by increased beta-cell hyperplasia (both from a increase in beta-cell replication and neogenesis) as well as beta-cell hypertrophy (i.e., an increase in beta-cell size) [1]. However, it is unclear what are the key growth factors and/or nutrients which promote this speciﬁc increase in beta-cell mass.

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Pancreatic Beta-Cell Death Introduction The net change in beta-cell mass is accounted for by the degree of beta-cell growth (i.e., the sum of the extent of replication, neogenesis and cell size) minus the degree of beta-cell death. In some instances, as in the pathogenesis of type 2 diabetes (see later), there is a measurable increase in beta-cell growth, but this is outweighed by the incidence of beta-cell death and as a result the net beta-cell mass is reduced [1,9]. In contrast, there does appear to be some islet cell apoptosis during the rapid increase in beta-cell mass neonatally, but this is overshadowed by the burst of beta-cell growth (Fig. 1A) [14]. Thus, beta-cell death is an important component when considering the regulation of beta-cell mass.

Types of Cell Death There are likely at least three ways in which a beta cell might die, either by necrosis, apoptosis or authophagic cell death. Necrosis has generally thought to be an uncontrolled form of cell death (usually a result of depleting adenosine triphosphate (ATP) levels to an extent where cell survival is irreversibly compromised) induced by toxins or physical insults [52]. However, necrosis can be caused by certain inﬂammatory cytokines (e.g., tumor necrosis factor (TNF)-alpha) to override apoptosis when it is inhibited during viral infection [53], or by DNA damage [54]. This has given rise to the term “programmed necrosis,” although it remains controversial and not universally accepted. Morphologically, a necrotic cell can be viewed as having a vacuolation of the cytosol, breakage of the plasma membrane and local inﬂammation likely induced by leakage of cellular contents. The nucleus of a necrotic cell can appear to be disﬁgured but does not show the nuclear condensation found in an apoptotic cell [52,55]. Apoptosis is also referred to as programmed cell death-1 (PCD-1) [56]. It is an intrinsic ATP-requiring mechanism allowing mammalian cells to commit suicide and is controlled by a variety of signaling pathways [52,57,58]. Apoptosis involves zymogen-like activation of certain members of a family of cysteine-proteases, known as caspases [57,58]. It is initiated by so-called death receptors and/or release of pro-apoptotic factors from mitochondria [58]. Morphologically, an apoptotic cell is characterized by nuclear condensation and DNA fragmentation, apparent shrinkage of the cytosolic compartment, but with no breach of the plasma membrane [52]. Apoptotic cells are removed by phagocytic cells without any notable local inﬂammatory response, which is unlike necrosis [52,56]. Autophagic cell death has recently been proposed

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as a probable third mechanism of cell death, also referred to programmed cell death-2 (PCD-2) [52,56,59–63]. It is found in certain human degenerative diseases [59–62], and is distinct from necrosis and apoptosis. Autophagy, under normal conditions, is an ATP-requiring regulated process in eukaryotic cells essential for cell survival that mitigates the degradation of protein complexes and organelles in their normal turnover [59–61]. However, if markedly upregulated, autophagy becomes a “double-edged sword” whereby it promotes cell death, independent of caspase activation [61,62]. Several autophagic genes (named ATGs) have been discovered that control the process of autophagy [61,64,65], but it remains unclear if upregulation of any of these genes can lead to autophagic cell death in a physiological setting. Indeed, although autophagic cell death does occur in some neurodegenerative diseases and cardiomyopathies, there is some debate as to whether it is part of a necrotic process, which also requires intracellular breakdown that may include autophagy [52,56]. Further experimental evidence is needed to ascertain whether autophagic beta-cell death might occur in the pathogenesis of diabetes.

Limitations of Measuring Beta-Cell Death Assessment of beta-cell death is a difﬁcult process that is technically limited. Although beta-cell necrosis and apoptosis can be distinguished in vitro [66,67], beta-cell death is hard to assess in an in vivo setting or from pancreatic pathological specimens unless examined by electron microscopy but this is rarely done. Most commonly, only apoptotic beta-cell death is measured in pancreatic sections [9,50,68], and necrosis and possible autophagic cell death are unaccounted. Moreover, apoptotic cells are efﬁciently cleared by macrophages in vivo. As such, the extent of beta-cell apoptosis, especially when analyzed in ex vivo pancreatic sections, is likely underappreciated.

Beta-Cell Death and the Pathogenesis of Diabetes Although mediated via distinct mechanisms, it is now widely acknowledged that both type 1 and type 2 diabetes are conceptually similar diseases of insulin secretory deﬁciency and decreased beta-cell mass [1,69]. The marked degree of beta-cell death that occurs in type 1 diabetes is mediated via a speciﬁc autoimmune and cytokine mediated attack on the beta cells that is well established, and details of which can be found in excellent reviews [3,4]. Type 2 diabetes is most commonly linked with obesity [70,71]. Currently in the United States about one third of obese human subjects acquire type 2 diabetes, due to a combination of insulin secretory dysfunction and a reduced beta-cell mass [1,5]. In obese or lean diabetic humans, as well as rodent

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models, the onset of type 2 diabetes is marked by a decrease in beta-cell mass that can no longer compensates for the metabolic load, especially insulin resistance associated with obesity. This beta-cell loss occurs from a marked increase in beta-cell apoptosis that far outweighs modest increases in the rate of beta-cell growth (Fig. 1A) [9,68,72–75]. In humans, the increased beta-cell apoptosis in the later stages of type 2 diabetes is further enhanced by the formation of islet amyloid deposits [76,77] (see the chapter by Clark). The rate of beta-cell loss varies between individuals, as does the onset of type 2 diabetes that is likely related to the extent of beta-cell loss [9]. If allowed to proceed untreated a “point of no return” for critical beta-cell mass is reached and a permanent state of type 2 diabetes sets in that requires insulin replacement therapy [78]. There is thought to be both a genetic and environmental component for the susceptibility of type 2 diabetes [5,79]. In addition, the “baseline beta-cell mass” in the adult that is essentially acquired from the critical burst of neonatal beta-cell growth may play a role (Fig. 1A) [1,14]. The lower the baseline beta-cell mass the smaller the capacity to compensate for an increase in metabolic load and/or insulin demand as found in pregnancy or obesity. As such, a low beta-cell mass could represent an increased risk of gestational diabetes, or progression to obesity-linked type 2 diabetes [1,80]. This may account for the increased incidence of type 2 diabetes associated with low birth weight [81]. However, key to the onset of type 2 diabetes are certain factors that may trigger the increased rate of beta-cell death. Several such factors have been proposed, often referred to as “beta-cell stresses,” which are outlined below (Fig. 2). Many of these beta-cell stresses have been shown to induce beta-cell apoptosis in in vitro studies, but it is so far unclear whether any of these have individual pathophysiological relevance to the onset of obesity-linked type 2 diabetes in vivo. In all likelihood it will be a combination of these “stress factors.”

Development of Beta-Cell Stress in the Pathogenesis of Type 2 Diabetes As obesity develops, there is ever increasing hyperlipidemia and lipid deposition in tissues other than adipocytes including liver, muscle, and islet beta cells [82–85]. This abnormal accumulation of lipid, particularly in muscle and liver, is a major contributor to the development of insulin resistance, which adds to the elevated metabolic load. Also, adipocyte-derived cytokines increase as the fat mass increases [83,86,87], which together with additional local inﬂammation further contribute to increasing insulin resistance. The accumulating insulin resistance and elevated metabolic load place an increased compensatory demand on the pancreatic beta-cell to produce and secret more insulin [79,88]. Chronically, insulin secretory dysfunction and

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OBESITY Adipocyte derived cytokines

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Fig. 2. Stresses contributing to beta-cell death in the pathogenesis of obesity-linked type 2 diabetes. Ongoing obesity increases circulating lipids and adipocyte-derived cytokines that contribute to insulin resistance and an increased metabolic load. In the latter stages, this is added to by increasing ﬂuctuations in circulating glucose as beta-cell dysfunction arises. This results in various stresses on the beta cell (outlined in detail in the text) that can lead to beta-cell apoptosis and/or necrosis thereby reducing the beta-cell mass that marks the onset of obesity-linked type 2 diabetes

loss of beta-cell mass begin to arise together with developing hyperglycemia [1]. The hyperglycemia places additional demand on the already hard-working beta-cell. With time beta-cell stresses develop which contribute to the induction of beta-cell apoptosis, thereby reducing beta-cell mass and leading to the onset of type 2 diabetes. Some of these beta-cell stresses are outlined below. Endoplasmic Reticulum (ER) Stress ER stress is linked to the unfolded protein response (UPR) that is generally a survival mechanism where by malformed and/or misfolded proteins forming in the ER set off a response that leads to a downregulation of general protein synthesis [89,90]. However, if the UPR is strong or extended chronically this can induce ER stress and apoptosis [89,90]. Pancreatic beta cells appear to be unusually susceptible to ER stress [90,91]. In transgenic mouse models where key regulatory element genes of the UPR have been knocked out (e.g., protein endoplasmic reticulum kinase (PERK) or p58IPK [92,93]) mild diabetes develops because of a selective increase in beta-cell apoptosis leading to a reduction in beta-cell mass [89,91]. However, some care needs to be taken

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with the interpretation of these mouse models since they also deﬁcient in adipocyte fat stores (W. Landiges and M. Katze, personal communication [92]). CHOP, which lies downstream of PERK and p58IPK in the UPR signaling pathway [91], is also a key element in regulating adipocyte differentiation [94,95]. As such, the decrease in beta-cell mass found in PERK-null mice could be secondary to a failure to appropriately store lipid in adipocytes and accompanying hyperlipidemia. Nonetheless, another mouse model, the Akita mouse [96], acquires spontaneous diabetes because of a mutation in the preproinsulin-2 gene (Cys96Tyr) that renders a malformation of proinsulin-2 in the ER because it cannot form the appropriate disulﬁde linkage within proinsulin-2 [96]. As a consequence, marked beta-cell ER-stress arises resulting in apoptosis and reduced beta-cell mass to cause progressive hyperglycemia and diabetes, not associated with insulitis or obesity [56]. These, and certain other genetic-based models [97–100] illustrate that chronic ER stress can lead to a diabetic state via chronic activation of the UPR [90,91]. But they are uncommon, and somewhat removed from the pathophysiology of regular type 2 diabetes. It has been shown that certain fatty acids [101], cytokine-induced production of NO [102], and depletion of ER [Ca2+] can induce ER stress in beta cells that may contribute to beta-cell loss in the pathogenesis of type 2 diabetes [103], although this has yet to be demonstrated in vivo. In addition, it is possible that the marked increased demand for insulin production in a hard working beta-cell placed in an obese/insulin resistant setting, perhaps additionally mediated via extended hyperglycemic excursions, could also lead to chronic ER stress and induce beta-cell apoptosis. This, in turn, would also contribute to the reduced beta-cell mass that marks the onset of type 2 diabetes. Amyloid Stress In human type 2 diabetes there is acquired islet amyloid deposition that also contributes to the reduction in beta-cell mass [77]. For further details please refer to Clark’s chapter. The major component of this amyloid is a small protein called islet amyloid polypeptide (IAPP) [104]. ProIAPP is normally synthesized and processed in parallel with proinsulin, and normally located in the lumen of an insulin secretory granule. ProIAPP synthesis is upregulated in parallel to proinsulin [105] and can accumulate to form oligomeric ﬁbrils. IAPP oligomers have been shown to induce beta-cell apoptosis [106], perhaps in part by induction of ER stress. The accumulation of IAPP ﬁbrils in an obese/insulin resistant prediabetic setting could contribute to the reduction in beta cell mass that leads to the onset of type 2 diabetes. Lipid Stress There is hyperlipidemia in the obese/insulin resistant setting, and lipid deposits are abnormally found in many tissues other than adipocytes, including

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pancreatic beta cells [107]. Chronic accumulation of lipid in beta cells causes beta-cell dysfunction and apoptosis, a phenomenon often referred to as lipotoxicity [107]. It is thought that long-chain acyl-CoA moieties and local ceramide production are the most likely initiators of beta-cell lipotoxicity [107,108], but the mechanism as to how they actually promote beta-cell apoptosis is currently unclear. Indeed, this may occur by several means. Certain fatty acids have been shown to induce beta-cell ER stress [101]. It has been proposed that long-chain acyl-CoA induces the activation of novel protein kinase-C (PKC) isoforms that then Ser/Thr phosphorylate certain proteins that are key to beta-cell survival (e.g., IRS-2) leading to there accelerated degradation and consequently making the beta-cell more susceptible to apoptosis [1,109]. Likewise fatty acid accumulation may also interfere with the activation of protein kinase B (Akt/PKB), another key survival protein for beta cells [1,80,110], leading to increased vulnerability for beta-cell apoptosis. Chronic exposure of beta cells to fatty acids can also interfere with mitochondrial activity [111,112], which in turn compromises beta cells’ ability to appropriately generate secondary signals for normal function as well as lead to irregular accumulation of intracellular [Ca2+]i to cytotoxic levels [112]. Fatty acids can also adversely alter gene expression patterns in beta cells, particularly of key transcription factors, that eventually leads to beta-cell dysfunction and death [113]. It should also be noted that in the later stages of the pathogenesis of type 2 diabetes, when there are increased excursions in postprandial glucose [5], hyperglycemia works synergistically with hyperlipidemia, accelerating the incidence of beta-cell death and onset of type 2 diabetes. This has been termed glucolipotoxicity [114]. Oxidative Stress The beta cell is somewhat unique in that its metabolism is not only geared for generating ATP, but also secondary signals that regulate normal beta-cell functions [115,116]. In an obese/insulin resistant environment, the increased metabolic load places an increased demand on the beta cell that enhances its metabolism. In the pathogenesis of obesity-linked type 2 diabetes, this is initially driven by the hyperlipidemia and then augmented late by the hyperglycemia. The latter is often referred to as glucotoxicity. A byproduct of increased oxidative metabolism are reactive oxygen species (ROS), that if not rapidly deposed of intracellularly become cytotoxic [117]. Intriguingly, the beta cell has unusually low levels of antioxidant enzymes, especially compared to hepatocytes [118]. As such, ROS are more likely to increase in beta cells and cause oxidative damage leading to apoptosis [118,119]. Pharmacological or genetic strategies to increase antioxidant enzymes in beta cells have shown a degree of protection against beta-cell apoptosis [119].

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Metabolic Stress Increased beta-cell metabolism, whether due to chronically increased lipid and/or glucose in a worsening obese/insulin resistant setting, might also have other detrimental effects on beta-cell function and survival. For example, chronically increased beta-cell mitochondrial metabolism could generate unusually high levels of secondary signals, such as [Ca2+]i and [cyclic AMP]i, in addition to ROS, that if not appropriately buffered will accumulate to abnormal cytotoxic levels and also contribute to increased beta-cell apoptosis and necrosis [120,121]. Also, certain nutrient sensing regulatory enzymes might be chronically activated in a hyperglycemic environment, such as the Ser/Thr protein kinase mTOR [36]. Chronic activation of mTOR by glucose and other nutrients can lead to degradation and downregulation of key survival factors in beta cells, including IRS-2 and Akt/PKB, and this then leads to increased beta-cell apoptosis [86,122]. Inflammatory Stress It has become apparent that the obese/insulin resistant state is also an inﬂammatory one [24,69]. Various cytokines are released from adipocytes, and because of the increased adipocyte mass in obesity there is a corresponding increase in circulating adipocyte-derived cytokines [83,85–87]. There are also increased numbers of macrophages in obesity and this leads to an increase in cytokine production [85]. Moreover, it has been indicated that elevated glucose can increase cytokine production within pancreatic islets [123], although whether this is from islet cells or local macrophages is unclear. Nonetheless, there is an increase in cytotoxic cytokines (e.g., TNF-alpha, interleukin (IL)-6, IL-1beta and others [83,85]) that can promote beta-cell apoptosis in an obese/insulin resistant state. Cytokines can activate various signaling pathways in the beta-cell leading to apoptosis (as described below (see section on Signal Transduction Pathways below), which would contribute to decreased beta-cell mass for the onset of obesity-linked type 2 diabetes. Chronic Autophagic Stress Under normal circumstances, insulin secreted from the beta cell is rapidly replenished by a corresponding increase in proinsulin biosynthesis and processing [124]. However, insulin secretion and production are independently regulated in the beta cell [124]. Under some circumstances when insulin secretion is inhibited, insulin production is upregulated, but then there is also increased intracellular degradation of insulin stores by autophagy [125–128]. As such, the insulin secretory granule stores in a beta cell are mostly maintained at optimal levels [126]. However, in an obese/insulin resistant setting where there is acquired insulin secretory dysfunction [5,79], insulin production can be elevated in a effort to compensate for the increased metabolic

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demand [129]. As such, an imbalance between insulin secretion and production arises and there is increased autophagy to maintain insulin stores [127,128]. However, autophagy is a “double edged sword” and if allowed to go on chronically may lead to autophagic cell death [61]. Thus, in a beta cell working hard to produce insulin yet defective in insulin secretion, as may occur in an obese/insulin resistant situation, one can envisage that authophagic beta-cell death can also contribute to the loss of beta cells in the pathogenesis of type 2 diabetes.

Signal Transduction Pathways for Control of Beta-Cell Growth and Death The study of signal transduction pathways that control pancreatic beta-cell growth and death is in its relative infancy and consequently little is known in detail. However, some elements in certain growth factor signaling pathways have been found to be important for beta-cell growth and survival (Fig. 3A), as have certain elements in cytokine signaling pathways for beta-cell death (Fig. 3B). These are brieﬂy described below.

Signal Transduction for Beta-Cell Growth The best-characterized growth factor signal transduction pathways that can mediate beta-cell growth and survival are those activated by IGF-1, growth hormone, and prolactin. Growth hormone and prolactin activate similar signaling pathways [130]. Growth hormone or prolactin, on binding to their respective receptors, causes dimerization of the receptor that in turn associates and activates the tyrosine protein kinase, Janus kinase-2 (JAK-2) [130]. JAK-2 then predominately phosphorylates signal transducer and activator of transcription-5 (STAT5) which then migrates to the nucleus to activate certain factors that promote beta-cell replication (Fig. 3A), including preadipocyte factor-1 (Pref-1) [130] and cyclin-D2 [131]. STAT5 also induces the expression of the anti-apoptotic protein Bcl-xL and members of the SOCS (suppressors of cytokine signaling) family [130,132], that could also play a role in promoting beta-cell survival. IGF-1 binding to the IGF-1 receptor on a beta cell activates an intrinsic tyrosine kinase activity that then phosphorylates adaptor molecules such as SH2domain containing protein (Shc) and insulin receptor substrate 2 (IRS2) [2,133] (Fig. 3B). IRS2 is the predominant IRS-family member expressed in beta cells [1,134]. Tyrosine phosphorylation sites on such adaptors act as speciﬁc docking sites for activation of certain signaling proteins that bind via their SH-2 domains [135]. The growth factor receptor binding protein-2

A IGF-1 GH IGF-1 Receptor

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Fig. 3A,B. Signal transduction pathways that contribute to beta-cell growth and death. Several signaling pathways can contribute to beta-cell growth and death. For beta-cell growth (A), IRS 2 signaling (as may be activated by IGF-1), and JAK-2/STAT5 signaling as activated by growth hormone (or prolactin) are the better characterized and depicted here. For beta-cell death (B), inﬂammatory cytokine signaling pathways (as activated by interleukin (IL)-1beta, tumor necrosis factor (TNF)-alpha, IL-6, and/or interferon (IFN)gamma) are best characterized and depicted here. Details of these signal transduction pathways are described in the text

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(Grb-2)/mammalian Son of Sevenless (mSOS) complex associates with speciﬁc tyrosine phosphorylated sites on Shc and IRS-2 in beta cells [2,133,136]. The mSOS has guanine nucleotide exchange (GEF) activity for small GTP-binding protein, Ras. Once the Grb-2/mSOS complex associates with Shc and/or IRS-2, mSOS is activated and loads Ras with GTP for it to become activated. The Ser/ Thr protein kinase Raf-1 then speciﬁcally associates with activated Ras and as a consequence in then activated. This is followed by Raf-1 mediated phosphorylation activation the MAP kinase kinase, MEK-1, that then phosphorylates the MAP kinases, extracellular-signal regulated kinase (Erk)-1 and -2, for their activation. Activated Erk-1 and -2 can then phosphorylate downstream protein kinases for their activation (e.g., p90 ribosomal S-protein kinase—p90RSK), or migrate to the nucleus to promote transcriptional activation of beta cell genes (including the insulin gene [137]) that promote growth and survival [2]. Tyrosine phosphorylation of IRS-2 also leads to binding of the p85 regulatory subunit of phosphatidylinositol 3′-kinase (PI3K) (at alternative IRS-2 phosphotyrosine sites to that for Grb2/mSOS complex [135,136]) that in turn activates the p110 catalytic subunit of PI3K. Increased PI3K activity generates PtdIns(3,4,5)P3 from PtdIns(4,5)P2 in the plasma membrane that act as docking sites for the PH-domain of Akt/PKB [80,133]. Plasma membrane located phosphoinositide-dependent protein kinase (PDK-1) then phosphorylates Akt/PKB at Thr308 for partial activation and subsequent phosphorylation at Ser473 (perhaps by target of rapamycin complex TORC2 [138,139] and/or autophosphorylation [140]) results in full Akt/PKB activation. The PI3K-mediated increase in PtdIns(3,4,5)P3 may also activate certain PKC isoforms including the atypical PKCzeta [141,142]. Akt/PKB has a vast array of protein phosphorylation substrates [80,143]. These include glycogen synthase kinase 3 (GSK3), and the transcription factor Foxo1, which are essentially inhibited by Akt/PKB phosphorylation. Downstream of GSK inactivation, the transcription factor betacatenin is negatively regulated whereas cyclin-D is stabilized in the nucleus. In contrast, the protein kinase complex TORC1, anti-apoptotic proteins murine double minute protein-2 (Mdm2) and BAD, the transcription factors CREB (cyclic AMP response element binding protein) and Rb (retinoblastoma protein), and the cell cycle regulatory protein p21CIP are selected protein substrates essentially activated or stabilized by Akt/PKB phosphorylation [80,143– 145]. Downstream of Mdm2 phosphorylation, the transcription factor p53 is negatively controlled [145,146]. Bad phosphorylation prevents apoptotic pore formation in mitochondria. Collectively, the result of these downstream effects of Akt/PKB activation is to promote beta-cell growth and survival [147]. In addition, Akt/PKB mediated phosphorylation of the TORC1 complex leads to phosphorylation activation of p70S6K and 4E-BP1 that increases general protein synthesis at the translational level [36], and likely contributes to increasing beta-cell size [2,147].

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Certain nutrients, including glucose, can also activate some downstream elements in IRS-signaling pathways in islet beta cells, independent of IRS2. Glucose can activate Erk-1 and -2 via inducing an upstream Ca2+dependent activation of protein kinase A (PKA) [148,149] and/or a novel PKC isoform [109]. Glucose can also activate TORC1 and downstream p70S6K in a manner independent of Ca2+ and cyclic AMP, probably via mammalian target of rapamycin (mTOR) acting as an ATP/metabolic sensor [36,148]. It has also been shown that glucose and GLP-1 independently control IRS2 gene expression levels, which in turn lead to activation of IRS2/PI3K/Akt/PKB signaling [150,151]. This may be one means by which glucose can control beta-cell growth and survival. Indeed, it should be noted from transgenic mouse gene knockout studies that IRS2 signaling, via PDK1/PI3K/PKB is critical for beta-cell growth and particularly survival [134,152–155]. Moreover, speciﬁcally increasing IRS2 and Akt/PKB in beta cells has been shown to be protective to beta cells and delay the onset of diabetes in certain mouse models [153,156]. However, the therapeutic potential of increasing Akt/PKB activity has been questioned because of potential oncogenic effects [80,155], but since IRS2 is rapidly turning over in beta cells, regulation of IRS2 expression to promote beta-cell survival may warrant some therapeutic merit [150].

Signal Transduction for Beta-Cell Death The signaling pathway that leads to ER stress induced apoptosis has been relatively well deﬁned in other cell types (see reviews [90,91] for details), and is presumed to be similar in beta cells. However, little is known in detail about the signaling mechanisms that cause beta-cell apoptosis via changes in betacell metabolism, although downstream activation of stress MAP kinases, cJun kinase (Jnk)-1/-2 and p38 may be involved [1]. For cytokine-mediated inﬂammatory stress the signaling pathway leading to apoptosis will depend on the speciﬁc cytokine (Fig. 3B). For beta-cell death in type 2 diabetes, IL-1beta, TNF-alpha and IL-6 signaling are likely most relevant, whereas in cytokine mediated beta-cell destruction in type 1 diabetes IL-1beta and interferon (IFN)-gamma are relevant [69]. Interlukin-1beta on binding to and activating its receptor leads to an association of the adaptor molecule myeloid differentiation factor-88 (MyD88) (Fig. 3B). MyD88, via its “death domain,” associates with and activates the Ser/Thr protein kinase IRAK (IL-1 receptor-associated kinase). Then the adaptor molecule, tumor necrosis factor receptor-associated factor (TRAF)-6, is activated that via another adaptor, evolutionarily conserved signaling

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intermediate in Toll pathways (ECSIT), leads to activation of the mitogen activated kinase (MAPK) kinase, MEKK-1, to phosphorylate and activate the stress MAPKs, JNK-1/-2, and p38 [69,157]. This leads to activation of transcription factors such as activator protein-1 that contribute to transcribing genes needed to induce the apoptotic program [157,158]. Activation of TRAF6, via another adaptor protein, TAB (TAK-1-binding protein), also leads to activation of the Ser/Thr protein kinase, TAK-1 (transforming growth factorbeta activating kinase), which then activates another kinase, IkappaK (IkappaB protein kinase) [69,158,159]. IkappaK phosphorylates the protein IkappaB releasing it from association with the transcription factor, nuclear factor (NF)-kappaB. The IkappaB (inhibitor of NF-kappaB) is then ubiquitinated and degraded. The liberated NF-kappaB translocates to the nucleus where it can induce gene expression needed for the apoptotic program [157,159]. Interleukin-1 beta receptor activation may also lead to PKCdelta activation that may also contribute to beta-cell apoptosis [69]. In beta cells, TNF-alpha is thought to signal through the TNF-alpha receptor-1 (TNF-alphaR1) [69]. The activated TNF-alphaR1 then associates with the adaptor TNFalpha-receptor1-associated death domain protein (TRADD) via its “death domains” (Fig. 3B). Another adaptor, Fas-associated death domain protein (FADD), is recruited along with the other adaptor protein TRAF2 and death-domain kinase, RIP. Via interaction with TRAF2 the Ser/Thr protein kinase NIK (Nck interacting protein kinase) is activated, leading to downstream phosphorylation and activation of IkappaK. The, IkappaK mediated phosphorylation of IkappaB leads to NF-kappaB translocation to the nucleus and transcription of proapoptotic genes [159,160]. Tumor necrosis factor-alpha may also signal via TNF-alpha receptor-2 (TNFalphaR2). The activated TNF-alphaR2, via its death domain, then recruits the adaptors TRAF1 and TRAF2 that, via association and activation of apoptosis signal-regulating kinase-1 (ASK-1), leads to sustained activation of the stress MAPKs, JNK-1/-2, and p38 [161], and subsequent apoptosis. Interleukin-6 on binding to its receptor leads to JAK-2 activation and STAT3 phosphorylation activation. Unlike growth hormone or prolactin activation of STAT5, STAT3 activation leads to transcription of genes that can promote apoptosis [162]. Likewise, IFN-gamma signaling is via JAK-2/ STAT1 activation leading to proapoptotic gene transcription. Among the genes that STAT1 and STAT3 activate are members of the SOCS family, which feedback to suppress cytokine signaling. However, in chronic inﬂammatory stress, SOCS may have detrimental effects on cells. It has been shown that SOCS-1 and -3 can bind to IRS1 and -2 to cause their ubiquitination and subsequent degradation, that in beta cells would in turn jeopardize survival [1,163,164].

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Summary and Closing Remarks Although under normal conditions in the adult the pancreatic beta-cell mass might appear to be relatively static and beta-cell turnover very low [16], it has a remarkable capacity to adapt to increases in the metabolic load and can increase in order to meet the increased demand for insulin. Failure to do this results in type 2 diabetes. As such, type 2 diabetes, like type 1, is a disease of relative insulin insufﬁciency. The onset of type 2 diabetes is associated with decreased beta-cell mass due to a marked increase in beta-cell apoptosis. This is likely caused by several “beta-cell stresses” that chronically occur in an obese/insulin resistant environment. It is unlikely that any one of these stresses, in itself, is detrimental to the beta cell, but collectively they could be catastrophic, signiﬁcantly reducing beta-cell mass. As such, in considering novel therapeutic strategies to delay the onset of type 2 diabetes, eliminating the stress on the beta cell and/or protecting and promoting beta-cell survival hold merit. There has been an idea put forward that a pharmacological means of promoting beta-cell regeneration could also have therapeutic potential to delay the onset of diabetes. However, this would be relatively futile unless the ongoing beta-cell death is also stopped. There is much interest in examining the signal transduction pathways that control beta-cell growth and death with the goal of identifying targets for agents promoting beta-cell survival. In this regard, it should be noted that these signal transduction pathways are either compromised (e.g., insulin/IRS signaling) or activated (e.g., inﬂammatory IL-1beta, TNF-alpha, and IL-6 signaling) in liver, muscle, and fat in obesity and are major contributors to the insulin resistant state [1,82,83,86,87,165]. As such, a therapeutic strategy designed for a controlled upregulation of IRS2 signaling and/or downregulation of IL-1beta/TNF-alpha/IL-6 signaling in beta cells should also be advantageous for reducing insulin resistance in vivo. Moreover, alleviating insulin resistance would be of further beneﬁt to the beta cell by reducing the metabolic load and hence demand for insulin. It is anticipated that in the next few years such therapeutic strategies will arise and it will be interesting to see if they are effective at delaying or even preventing the onset of type 2 diabetes. Considering that cytokine mediated attack of beta cells is important in the pathogenesis of type 1 diabetes, some of these approaches may also be applicable to treat this form of the disease as well. Acknowledgments. It is realized that this chapter is a brief overview of the regulation of beta-cell growth and survival, and many details, and excellent work, from other researchers has been omitted. Unfortunately, there is only so much space one can ﬁll, and my apologies go out to those

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whose work I may have overlooked or details I have not sufﬁciently described. Some of the ideas and work outlined here have in part been possible through grants provided by the National Institutes of Health, DK 50610 and DK55267.

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13. Beta-Cell Replication Seth J. Salpeter and Yuval Dor

Summary. Patients suffering from type 1 and type 2 diabetes exhibit a decrease in the mass of insulin-producing beta cells. Both the ability to generate and expand large amounts of transplantable beta cells and the capacity to encourage beta-cell proliferation in the patient represent potential cures for the disease. Understanding the basic cell cycle machinery responsible for the replication of pancreatic beta cells is therefore an important challenge in diabetes research today, in hopes that it will provide useful insights into betacell growth and proliferation. Though for many years pancreas biologists believed that adult beta cells emerged from progenitor cells and remained post-mitotic throughout their lifetimes, recent work has demonstrated that adult beta cells are a dynamic and replicating population. In light of this new understanding of pancreatic beta cells, much attention is currently being focused on the regulation of the replication process. However, even as biologists focus on the particular machinery involved in division, a proper understanding can only be obtained in light of beta-cell development, origins, and dynamics. In this review, we present a brief introduction to beta-cell development and origins, followed by a description of beta-cell proliferation machinery. We conclude with a discussion of a possible regulatory model for beta-cell proliferation.

Introduction The mechanisms involved in beta-cell replication pose a signiﬁcant scientiﬁc question on several fronts. One pressing cause for investigation lies in the necessity to improve therapy for type 1 and type 2 diabetes, diseases which Department of Cellular Biochemistry and Human Genetics, The Hebrew University– Hadassah Medical School, Jerusalem 91120, Israel

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afﬂict more than 200 million people worldwide. In type 1 diabetes, an autoimmune response leads to the destruction of beta cells. Type 2 diabetes, accounting for >90% of cases of diabetes, is typically associated with peripheral insulin resistance. However, recent evidence indicates that in addition to insulin resistance, this disease is associated with defects in beta-cell function as well as a loss of as much as 50% of beta-cell mass [1,2]. Thus, the possibility of regenerating or replacing beta cells offers immense therapeutic potential in both types of diabetes. Indeed, a clinical procedure for islet transplantation recently developed at the University of Alberta (the Edmonton protocol) offers for the ﬁrst time a satisfying cure for diabetes [3]. However, a major limitation of the Edmonton protocol is the severe shortage of donor islets. As a result of this problem, signiﬁcant efforts are directed towards the development of strategies for enhancement of beta-cell mass, in vivo (regenerative therapy) or in vitro for transplantation (cell therapy). Such efforts will greatly beneﬁt from a better understanding of the normal process by which beta cells replicate and factors that increase beta-cell replication [4]. The analysis of beta-cell replication is the subject of this chapter. Beyond clinical importance, understanding the mechanism of beta-cell replication represents a basic challenge for cellular biologists. Though long considered static, islets of Langerhans (composed mainly of beta cells and constituting the endocrine pancreas) are now appreciated as an active and developing organ that maintains the ability to respond to external stimuli [5] and continues to grow throughout life [6,7]. What is the basic mechanism by which the cells replicate? What are the factors involved in triggering beta-cell replication? Is there a population of beta cells with a greater potential for replication—or possibly even adult stem cells? We start by providing a brief overview of the possible origins of new beta cells during normal adult life in order to give perspective on the role beta-cell replication plays in overall beta-cell mass. We then describe some highlights of the molecular machinery for adult beta-cell proliferation and its regulation, and conclude by suggesting a model for the basic cell cycle replication machinery.

The Origins of Adult Beta Cells Progenitor Cells and Neogenesis Postnatal beta cells in rodents and humans can proliferate, even though at a low rate, as demonstrated by the incorporation of BrdU or [3H]thymidine [8–11]. This however does not necessarily mean that beta-cell replication is responsible for the generation of new beta cells: beta cells could proliferate at a low rate, but still be derived mainly from stem cells. Over the years, many different cells have been proposed as the source of new beta cells.

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For example, pancreatic ducts are considered by many a pool of adult stem cells constantly replenishing beta cells. This view is largely based on the fact that during embryonic development of the pancreas, primitive ducts give rise to Ngn3+ endocrine progenitors, which in turn differentiate to hormone producing cells including beta cells. In addition, static histologic observations are often interpreted as showing islets “budding” from adjacent ducts [12–15]. Similar to the hypothesis of ductal contribution to beta cells, it has been suggested that acinar cells may undergo transdifferentiation and convert into beta cells [16,17]. Other recent studies suggest that bone marrow cells [18,19], spleen cells [20], or intra-islet progenitor cells expressing nestin [21] may supply the growing beta-cell mass. Many of these ﬁndings have proved difﬁcult to reproduce [22–29], and so far the progenitor cell origins of beta cells have not been demonstrated by robust lineage tracing experiments.

Pre-Existing Beta Cells In light of these conﬂicting reports, a study was undertaken by Dor and colleagues to apply inducible lineage analysis to the adult beta-cell using a new method termed “genetic pulse-chase” [30]. Transgenic mice were generated in which tamoxifen-dependent Cre recombinase was placed downstream of the rat insulin promoter. These mice were then crossed with a reporter transgenic strain expressing human placental alkaline phosphatase (HPAP) in cells that undergo Cre-mediated recombination. Adult mice were injected with tamoxifen to label existing, differentiated beta cells (“pulse”), and the fate of these labeled beta cells was followed over time (“chase”). If beta cells are derived from pre-existing beta cells, the fraction of labeled beta cells should remain stable. If however dying beta cells are replaced by new beta cells derived from any type of stem cell (hence unlabeled), the fraction of labeled beta cells should decline with time. Strikingly, the fraction of labeled beta cells did not decline even a year after the original labeling. This led the authors to conclude that during adult life, new beta cells are largely the product of betacell proliferation rather than stem cell differentiation. Similar experiments were performed with mice following partial pancreatectomy, a procedure known to result in some regeneration of beta cells. Here again, the results demonstrated that most new beta cells were the progeny of beta cells that existed prior to the operation. Importantly, this experimental system was designed to determine the major source of new beta cells during postnatal life. Stem cells could still exist and give rise to small numbers of beta cells. Such stem cells could be recruited to action following speciﬁc injury conditions. These are important considerations because even a small contribution

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of stem cells could in principle have signiﬁcant biotechnological/therapeutic utility [4]. Other studies provide indirect support for the notion that beta-cell proliferation is a dominant process in beta-cell dynamics. For example, mice lacking the cell cycle proteins cyclinD2 or CDK4 show a progressive failure of beta-cell mass [31–33]. Since these genes are mainly expressed in differentiated beta cells, this supports the primacy of beta-cell proliferation. In addition, forced expression of the cyclin kinase inhibitor p27 in beta cells resulted in a progressive decline in beta-cell mass, again suggesting that beta-cell proliferation is essential for maintaining normal beta-cell mass [34].

Dedifferentiation of Beta Cells Interestingly, is has been suggested recently that differentiated beta cells can undergo a process of epithelial to mesenchymal transition giving rise to a proliferative population of insulin-negative cells capable in principle of redifferentiating into beta cells [35,36]. These studies are based on tissue culture studies using human islets. Lineage tracing experiments with cultured mouse islets, carried out in our group, have shown that beta cells can indeed dedifferentiate and lose expression of essential genes such as insulin, Pdx1, Glut2 and prohormone convertase. However, under normal culture conditions these dedifferentiated beta cells only rarely proliferate and are eventually eliminated from culture, leaving open the question of whether they can function as proliferating progenitors [37–39]. Notably, in vivo evidence suggests that under certain conditions, proliferating beta cells can undergo signiﬁcant phenotypic changes, including some that resemble epithelial to mesenchymal transition [40]. Understanding the extent and signiﬁcance of beta-cell dedifferentiation and epithelial to mesenchymal transition in vivo remains an important challenge.

Capacity of Beta-Cell Replication Given the understanding that beta-cell replication is indeed the primary factor in islet beta-cell expansion, the overall dynamics of beta-cell replication take on new importance. Before addressing speciﬁc pathways and replication machinery, we discuss normal beta-cell replication rates to shed light on the natural replication potential of beta cells in vivo. Early reports by Finegood and colleagues suggested that rat beta-cell proliferation rates are roughly 20% per day in pups, moving to 10% replication

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in adolescence and ﬁnally settling at around 2.0% in older rats [41,42]. Others found lower replication rates in older rats, suggesting the rate is roughly 0.8% [6] while more recent reports suggest that it may be as low as 0.07% [43]. Based on these calculations, various estimates of the life span of a betacell have emerged. Based on Finegood’s data, it was concluded that new beta cells live for 58 days while estimations of slower beta-cell turnover must assert that beta cells have much longer life spans. Human beta-cell replication rates has been shown to be signiﬁcantly lower than the murine rate thereby suggesting that in humans the beta-cell life span would have to be much longer [44]. Such reports raise the question of whether the slower beta-cell replication with age represents an inherent reduction in the beta cell’s capacity for replication or simply a lack of necessity for increased proliferation. One study treated 1-year-old mice with low doses of the beta-cell toxin streptozotocin to test the ability of older beta cells to re-enter the cell cycle [43]. Indeed, it was found that the beta-cell replication rate in this age group jumped from 0.08% per day to 0.93% per day during the two weeks post ablation period. Such studies suggest that even older beta cells retain the capacity for proliferation once a proper stimulus is provided.

Cell Cycle Machinery Before delving into the speciﬁc regulatory factors involved in beta-cell replication, a brief introduction is needed on general cell cycle progression. Some excellent recent reviews provide more in depth analysis of the topic [45,46]. Cyclin-dependent kinases (CDKs) and cyclins are generally responsible for driving forward the cell cycle from G1 to S then G2 and ﬁnally mitosis. Throughout this process, these activating factors are controlled by inhibitors (the CKI and INK families). Of special importance is the junction between G1 and S, where CDKs and cyclins are responsible for phosphorylating the Retinoblastoma (Rb) protein, which releases the E2F transcription factors and drives the cell cycle forward. When analyzing a speciﬁc tissue’s cell cycle regulation, it is important to keep in mind the variety of CDKs and cyclins in existence. To date, 9 different CDKs and 16 different cyclins have been identiﬁed. Speciﬁc tissues utilize a certain set of CDKs and cyclins while other tissues may not use these factors at all in their cell cycle progression. Furthermore, a tissue may have redundancy for any given factor. That is to say that in the absence of its principle CDK or cyclin, cells may make use of another cell cycle progression factor as a substitute. With these principles in mind we can now take a look at speciﬁc cell cycle elements within pancreatic beta cells.

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Cyclins Within the cyclin family there are three D-type cyclins: D1, D2, and D3. Each of these has the ability to form a complex with CDK4 or CDK6 and phosphorylate Rb. Speciﬁcally in beta cells, it has been found that cyclin D2 is the main cell cycle cyclin, with some evidence for expression of cyclin D1 [31,32,47,48]. The presence of cyclin D3 is controversial with some arguing it is not present [49] and others suggesting that there are observable levels of cyclin D3 mRNA and protein [50]. Expression levels aside, genetic studies have demonstrated that only cyclin D2 is necessary for normal pancreatic beta-cell development in the mouse. cyclin D1-null mice have normal islet number and size, while cyclin D2-null mice have decreased beta-cell mass and poor glucose regulation [31,32]. As mentioned above, an interesting aspect of the beta-cell replication machinery is the redundancy and ﬂexibility of the system. Though cyclin D2null mice have compromised beta-cell mass, there is a further reduction when one of the two cyclin D1 alleles is removed [32]. Another example of such redundancy comes from a study which overexpressed cyclin D1 in beta cells resulting in increased beta-cell proliferation [51]. Thus, despite having a smaller and possibly minimal role in beta-cell replication, beta cells can use cyclin D1 to compensate for a lack of cyclin D2 and can also employ it to achieve hyperplasia. Cyclin E and cyclin A have also been found within beta cells [47,52]. Generally, both cyclin A and cyclin E pair with CDK2 to drive the cell forward from G1 into S phase [53,54]. Despite noticeable expression levels, cyclin E-null mice have no defect in beta-cell development [55]. Possibly the most central question which remains to be solved is exactly how the cyclins are regulated (Fig.1). Several factors have been identiﬁed as possible regulators of cyclin D2 expression in beta cells. Mice overexpressing Akt in their beta cells were found to have an upregulation of cyclin D2 mRNA and protein [56], while another study found that activation of the Jak/Stat pathway increases cyclin D2 expression [57]. Some work has been done in other types of cells that may function to elucidate the beta-cell pathway. In mammary carcinomas, it was found that, the cyclin D1 promoter contains a STAT binding site that may function as a regulatory region [58]. In pituitary cells, the transcription factor Pitx2 was found to regulate cyclin D2 expression through a Wnt dependent signaling pathway [59]. Possibly the most interesting is the ﬁnding that in immune system B cells both the NF-kappa B pathway via C-myc and Mek1/2 via ERK directly regulate cyclin D2 transcription [60]. In contrast, repression of cyclin D2 gene transcription has been found

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to be mediated by Mad/Max complexes [61]. Further work on these pathways may be the key to uncovering the major factors involved in beta-cell replication. Beyond the speciﬁc pathway regulating cyclin D2 lies the question of whether it is regulated on the transcriptional or protein level. In mice overexpressing Akt, an increase in the amount of both cyclin D1 and cyclin D2 protein was found despite a decrease in the amount of cyclin D1 and D2 mRNAs [50]. Such a result indicates that cyclin controlled replication may be regulated at the translational or protein stability level and not by transcription. Paradoxically, despite tremendous efforts to understand the beta-cell replication machinery as a means of increasing islet mass for human therapy, little research has been done on cell cycle in human tissue. One study found low levels of cyclin D1 and CDK4 mRNA in human islets [48]. Further analysis of human islets will be necessary both to validate current mouse models and to further assist the efforts towards beta-cell therapy.

A: Signaling Pathways GH

Prolactin

B: Cell Cycle Machinery

GLP-1

Complex Formation: G0/G1

Insulin/IGF GHR

PRLR

GLP-1R IR

CDK4

IGF Glucose

CyclinD2

INK4A CIP/KIP P16,p18 P27/P21

Glut2 Irs-2 Jak2

PI3K

Inhibitors Released: G1/S Calcium

PDK-1

Stat-5

CDK4

Akt

CyclinD2 +

CDK2

Calcineurin S6K

CyclinA/E

NFAT FoxO1

Phosphorylation of RB, Release of E2F: S ?

Cell Cycle Reguation

pRB C-Myc

E2F

? Menin

DNA Replication

Fig. 1. Schematic of signaling pathways and molecular machinery of pancreatic beta-cell proliferation. See text for details

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Cyclin-Dependent Kinases Investigators have found that Cdk4 and Cdk2 are expressed in islets while Cdk6 expression is not found at all [47,49,52]. A groundbreaking study by Rane and colleagues [33] found that CDK4-null mice develop normal islets, however after weaning beta cells fail to undergo expansion and the mice ultimately develop hyperglycemia and diabetes. Ectopic expression of CDK4 speciﬁcally in islets of these mice rescues this phenotype indicating that betacell failure is the result of CDK4 deﬁciency speciﬁcally in beta cells (and not in progenitor cells) [49]. Importantly, expression of a constitutively active CDK4 that lacks a P16ink4a binding site results in dramatic islet hyperplasia, demonstrating that CDK4 is not only necessary but also sufﬁcient to drive beta-cell proliferation. This and more recent ﬁndings suggest that the INK4 inhibitors of CDK4 play an important role in restricting beta-cell proliferation [62]. Similar to cyclins, a pressing question exists regarding the regulation of CDK4 within beta cells. Fatrai and colleagues [50] examined this question in a transgenic mouse model where constitutively active Akt is expressed in beta cells and leads to islet hyperplasia. No increase in the amount of CDK4 protein was found, even though the presence of an intact CDK4 gene was absolutely essential for AKT-induced hyperplasia. Thus, there is no evidence at present that the expression of CDK4 is regulated during beta-cell proliferation. Similarly, Cozar-Castellano and colleagues [47] did not see any increase in the quantity of CDK4 in response to overexpression of hepatoycte growth factor or placental lactogen in islets. However, in a recent study of menin-deﬁcient mice (see below), Karnik and colleagues demonstrated that beta cells in such mice have increased CDK4 mRNA and protein levels, indicating a possible regulatory step at the level of CDK4 expression [63].

Cyclin-Dependent Kinase Inhibitors (CKI) INK4s The function of the INK4s is to directly suppress the activity of CDK4 by binding to the protein. The critical importance of these proteins in beta-cell proliferation was shown in studies on mice expressing an INK4-insensitive CDK4 mutant. These mice had a 10-fold increase in beta-cell mass [33]. Several other studies note the importance of other INKs within the beta-cell cycle. A recent study examined the role of p16 and showed that p16 upregulation is responsible for the observed decrease in beta-cell replication with age [62]. Furthermore, although p16ink4a knockout yields only a minor increase in beta-cell mass in older mice, p16ink4-deﬁcient mice show a more

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robust regenerative response to streptozotocin-mediated beta-cell damage compared with wild type mice. In another study, P18ink4c was shown to play a fundamental role in beta-cell control as knockout mice exhibit a 40% increase in beta-cell mass [64,65]. P15ink4b has also been found in adult islets in high levels [47,52,66]. Upregulating TGF-beta in the pancreas led to p15 overexpression which caused islet hypoplasia and diabetes [67]. Finally, combined knockout of both the p15 and p18 inhibitors causes mice to develop pancreatic endocrine tumors [66]. CIP/KIP Recent ﬁndings are beginning to illustrate the importance the CIP/KIP regulators p21, p27, and p57 in beta-cell regulation. Several studies have demonstrated high levels of p27 and p21 in mouse islets [34,47,63,68]. Early reports of mice lacking p27 throughout the body noted no striking phenotype of beta-cell hyperplasia [69,70]. However, a more recent report [34] shed new light on the important function of p27 in beta cells. Despite ﬁnding that p27 knockout in a regular beta-cell does not affect islet size, in mice lacking either the insulin receptor substrate 2 (Irs2-null) or the leptin receptor (Lepr-null), p27 deﬁciency was found to alleviate hyperglycemia and increase beta-cell mass. Furthermore, in mice over-expressing p27, beta-cell replication rates were signiﬁcantly decreased and islet hypoplasia ensued. This study suggests that p27 can be regulated by the PI3K pathway via Irs2. Despite this suggestion, it has been demonstrated in mice with activated AKT that there is no decrease in the overall amount of p27 protein [50]. Recently two groups undertook more speciﬁc analysis of p27 knockout mice [68,71]. Both studies noted the lack of p27 caused a two-fold increase in beta-cell mass. Despite this general agreement, Georgia and Bhushan [71] suggested that the increase of mass took place entirely in the embryonic stage and stopped during development, while Rachdi and colleagues [68] found an increase in beta-cell replication rate even at 3 months. Rachdi and colleagues further examined the signiﬁcance of p27 overexpression both in developing and adult mice and found that overexpression during development caused a drop in beta mass while in adult mice overexpression had no effect. P21Cip has been implicated as an important inhibitory factor and requires additional investigation. It has been found that p21 can be increased in beta cells in response to oxidative stress [72] and is upregulated in islets overexpressing AKT, Hepatocyte growth factor (HGF), and Placental lactogen (PL) [47,50]. However, p21-deﬁcient mice exhibit no defect in beta-cell proliferation or mass [73], suggesting that either p21 has no functional role in restricting beta-cell proliferation or that redundancy with other CKIs masks the effect A ﬁnal possible candidate for inhibiting beta-cell replication is p57. p57 is found in murine beta cells, though at signiﬁcantly lower levels than other

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factors [73]. Interestingly, human islet which express only minimal levels of p27 were found to express high level of p57 [74]. Importantly, studies of the focal form of persistent hyperinsulinemic hypoglycemia of infancy have implicated reduced expression of p57 as a potential cause for the excessive beta-cell proliferation [75]. No islet phenotype was reported for mice lacking p57 [76]. E2F and Rb The E2F family contains seven transcription factors that regulate the transition from G1 to S. Generally they are regulated by Rb which upon phosphorylation, releases the transcription factors and allows the cell cycle to progress. In islets, all seven E2Fs have been identiﬁed with 1, 4, and 6 being the most prominent [47,52,77,78]. Recently, the Rb gene was speciﬁcally deleted in beta cells [79]. Surprisingly, no effect on beta-cell mass or physiology was observed. Furthermore, Rb pocket protein homologues p107 and p130 were not upregulated, leading the authors to argue against the possibility that a similar protein replaced Rb within the cell cycle pathway. These results suggest, but do not prove, that Rb and pocket proteins may not play a signiﬁcant role in beta-cell cycle. In mice with E2F1 whole body knock-out there is lower beta-cell mass; however, the mice do not develop diabetes [80]. Principally, the islets of these mice seem to have impaired Pdx-1 expression. However, when both E2F1 and E2F2 are knocked out, mice have decreased beta-cell mass and develop diabetes [78]. Interestingly, despite the overall decrease in beta-cell mass in E2F1/E2F2 deﬁcient mice, there was an increase in beta-cell proliferation which was neutralized by increased apoptosis.

Cell Cycle Transduction Pathways A proper analysis of beta-cell replication requires a discussion of certain factors that may act immediately upstream of the classic cell cycle machinery. These proteins are the critical mediators that transduce mitogenic signals into activity within the cell cycle.

Phosphatidylinositol-3-Kinase (PI3K) Pathway A large body of evidence indicates that the insulin receptor PI3K pathway is central for the regulation of beta-cell proliferation and for determining ﬁnal beta-cell mass. In mice lacking the insulin receptor speciﬁcally in beta cells a mild decrease in beta-cell mass was observed [81,82], while a combined deﬁciency of IGF-1 receptor and insulin receptor in beta cells results in a signiﬁcant decrease in beta-cell mass at 2 weeks of age [83]. Additional studies examining other downstream factors within the PI3K pathway have yielded

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similar conclusions. In Irs-2 knockout mice a signiﬁcant drop is found in beta-cell mass yielding type 2 diabetes at 10 weeks [84,85]. Further downstream in the pathway, Pdk1 deﬁciency in beta cells results in a similar phenotype with a signiﬁcant decline in both beta-cell mass and beta-cell size [86], while beta cell-speciﬁc deletion of Pten results in islet hyperplasia [87]. The centrality of Akt activation in this pathway was demonstrated by the increase in beta-cell replication, mass and cell size in transgenic mice expressing a constitutively active Akt transgene in beta cells [56,88]. Despite this signiﬁcant ﬁnding, little progress has been made in understanding how AKT directly regulates beta-cell replication. One possibility lies in Akt’s regulation of Foxo1. Akt is known to phosphorylate Foxo1 causing nuclear exclusion and thereby preventing it’s anti-proliferative activity. When Foxo1 was made constitutively nuclear in a transgenic model, Pdx-1 expression was reduced and diabetes ensued [89]. Some groups suggest that nuclear Foxo1 inhibits proliferation by increasing p27 levels [90] and by reducing expression of cyclin D1 and cyclin D2 [91]. A more intriguing suggestion is that Foxo1 directly regulates Pdx-1 expression and the level of Pdx-1 regulates the expression of other beta-cell genes [92]. One study noted that Pdx-1 overexpression in islets of Irs2-deﬁcient mice was able to restore normal proliferation [93]. Furthermore, overexpression of Pdx-1 alone yields a signiﬁcant increase in beta-cell mass with double the number of replicating cells at 3 weeks of age. Further analysis is needed to better understand the role of Pdx-1 in beta-cell replication.

Calcineurin/Nuclear Factor of Activated T Cells (NFAT) Beyond the PI3K pathway, several other regulators have been found to play critical roles in beta-cell replication. A recent study on the signiﬁcance of calcineurin/NFATc demonstrated the signiﬁcance of calcium signaling on beta-cell proliferation [100]. Calcium is known to elevate calcineurin phosphatase leading to dephosphorylation of cytoplasmic NFATc followed by nuclear localization of NFAT. There, NFAT acts as an important transcriptional regulator. In mice lacking calcineurin phosphatase, NFAT is unable to enter the nucleus. The result of this mutation is a signiﬁcant decrease in betacell replication and mass. Conversely, when NFAT is constitutively active in the nucleus there is an increase in beta-cell replication and mass. This study highlights an important link between beta-cell physiology- manifested as ﬂuctuations in calcium levels- and beta-cell replication.

Menin The product of the Men1 gene, Menin, is a transcriptional regulator and tumor suppressor whose absence results in the MEN syndrome (multiple

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endocrine neoplasia). Beta-cell speciﬁc deletion of Menin in mice was found to cause dramatic beta-cell hyperplasia leading to insulinomas at 7 months of age [94,95]. A subsequent study demonstrated that lack of Menin causes a drop in p18 and p27 levels, while at the same time yielding an increase in the amount of CDK4 [63]. Furthermore, Menin was found to associate with the p27 and p18 promoters and methylates lysine 4 of histone 3 on these regions. Results from these studies point to the necessity of further investigation of epigenetic regulation in beta-cell dynamics.

c-Myc Finally, the product of the c-Myc gene appears to be a signiﬁcant regulator of pancreatic beta-cell proliferation. When c-Myc is overexpressed in beta cells, it causes all beta cells to enter the cell cycle within 24 h [96]. This dramatic increase in beta-cell replication precipitates a strong apoptotic response, thereby erasing existing beta-cell mass. However, when c-myc is overexpressed with Bcl-x (L), beta-cell apoptosis is prevented and insulinomas begin to form. Further studies of the mechanism of c-Myc activity indicated that cMyc overexpression leads to a dramatic increase in levels of cyclin D1, cyclin A, cyclin B1 and cyclin B2 [97]. Interestingly, c-Myc activation results in a signiﬁcant decrease in Pdx-1, Glut2 and insulin levels suggesting a potential coupling between beta-cell proliferation and dedifferentiation.

Extracellular Growth Factors Ultimately beta-cell replication is thought to be triggered by mitogens that lie upstream of intracellular transduction pathways. Numerous factors have been implicated as activators of beta-cell replication. These factors include prolactin, glucose, insulin, hepatocyte growth factor, epidermal growth factor, parathyroid hormone related protein, IGF-1, GLP-1, and others. The signaling pathways downstream of these factors that lead to beta-cell proliferation are not well understood, Furthermore, it is not known which of these factors is playing a signiﬁcant role in triggering beta-cell proliferation in the intact organism. Though this important issue is beyond the scope of this chapter, several excellent reviews address this topic [98,99].

Conclusion This review illustrates that beta-cell replication is a dynamic process drawing on numerous components to achieve regulation. The pathway starts with extracellular mitogens, is translated into intracellular mediators, which then affect cell cycle machinery composed of both activators and inhibitors.

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Altering a given step within these pathways may dramatically change the cell’s replicative potential. In terms of the basic cell cycle machinery, a simplistic model can be suggested based on our current data. According to this model, the beta cells’ decision to proliferate resides with CDK4 and cyclinD2. These proteins are regulated by the INK4A (mostly p16 and p18) and the CIP/KIP proteins (p27 and p21 in mice, p57 in humans). The removal of the inhibitors from the CDK4-CyclinD2 complex activates it and facilitates the phosphorylation of Rb or other members of the Rb family. Other mechanisms for activation of the CDK4–cyclinD2 complex, e.g., phosphorylation, are also plausible. It is also likely that CDK2 in tandem with either cyclin A or cyclin E plays a supplementary role in phosphorylating the Rb protein, downstream of CDK4 activity. Together the activation of CDK4 and CDK2 and subsequent phosphorylation of Rb causes the release of E2F and progression into S phase of cell cycle. When struggling with the basic questions of beta-cell cycle machinery, we must keep in mind that the vast majority of cell cycle research has been performed in vitro. Only recently have biologists started to address the molecular machinery of cell cycle progression in vivo. Furthermore, the mechanisms driving cell cycle progression in speciﬁc cell types such as beta cells are largely unexplored, and might prove quiet different from the generic cell cycle paradigm of cultured ﬁbroblasts. The pressing need for increasing beta-cell numbers as a therapy for beta-cell deﬁciency has catapulted diabetes researchers into the forefront of cell cycle biology. This tremendous challenge demands knowledge from several ﬁelds and, moreover, requires beta-cell biologists to push the boundaries of what is known in the ﬁeld of cell cycle.

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6. Montanya E, Nacher V, Biarnes M, Soler J (2000) Linear correlation between beta-cell mass and body weight throughout the lifespan in Lewis rats: role of beta-cell hyperplasia and hypertrophy. Diabetes 49(8):1341–1346 7. Skau M, Pakkenberg B, Buschard K, Bock T (2001) Linear correlation between the total islet mass and the volume-weighted mean islet volume. Diabetes 50(8): 1763–1770 8. Messier B, Leblond CP (1960) Cell proliferation and migration as revealed by radioautography after injection of thymidine-H3 into male rats and mice. Am J Anat 106:247–285 9. Meier JJ, Lin JC, Butler AE, Galasso R, Martinez DS, Butler PC (2006) Direct evidence of attempted beta cell regeneration in an 89-year-old patient with recent-onset type 1 diabetes. Diabetologia 49(8):1838–1844 10. Ritzel RA, Butler PC (2003) Replication increases beta-cell vulnerability to human islet amyloid polypeptide-induced apoptosis. Diabetes 52(7):1701–1708 11. Kassem SA, Ariel I, Thornton PS, Scheimberg I, Glaser B (2000) Beta-cell proliferation and apoptosis in the developing normal human pancreas and in hyperinsulinism of infancy. Diabetes 49(8):1325–1333 12. Kim SK, MacDonald RJ (2002) Signaling and transcriptional control of pancreatic organogenesis. Curr Opin Genet Dev 12(5):540–547 13. Bonner-Weir S, Baxter LA, Schuppin GT, Smith FE (1993) A second pathway for regeneration of adult exocrine and endocrine pancreas. A possible recapitulation of embryonic development. Diabetes 42(12):1715–1720 14. Bonner-Weir S, Toschi E, Inada A, Reitz P, Fonseca SY, Aye T, Sharma A (2004) The pancreatic ductal epithelium serves as a potential pool of progenitor cells. Pediatr Diabetes 5 Suppl 2:16–22 15. Bonner-Weir S, Sharma A (2002) Pancreatic stem cells. J Pathol 197(4):519–526 16. Lipsett M, Finegood DT (2002) beta-cell neogenesis during prolonged hyperglycemia in rats. Diabetes 51(6):1834–1841 17. Minami K, Okuno M, Miyawaki K, Okumachi A, Ishizaki K, Oyama K, Kawaguchi M, Ishizuka N, Iwanaga T, Seino S (2005) Lineage tracing and characterization of insulin-secreting cells generated from adult pancreatic acinar cells. Proc Natl Acad Sci USA 102(42):15116–15121 18. Ianus A, Holz GG, Theise ND, Hussain MA (2003) In vivo derivation of glucosecompetent pancreatic endocrine cells from bone marrow without evidence of cell fusion. J Clin Invest 111(6):843–850 19. Lee VM, Stoffel M (2003) Bone marrow: an extra-pancreatic hideout for the elusive pancreatic stem cell? J Clin Invest 111(6):799–801 20. Kodama S, Kuhtreiber W, Fujimura S, Dale EA, Faustman DL (2003) Islet regeneration during the reversal of autoimmune diabetes in NOD mice. Science 302(5648): 1223–1237 21. Lechner A, Leech CA, Abraham EJ, Nolan AL, Habener JF (2002) Nestin-positive progenitor cells derived from adult human pancreatic islets of Langerhans contain side population (SP) cells deﬁned by expression of the ABCG2 (BCRP1) ATP-binding cassette transporter. Biochem Biophys Res Commun 293(2):670–674 22. Suri A, Calderon B, Esparza TJ, Frederick K, Bittner P, Unanue ER (2006) Immunological reversal of autoimmune diabetes without hematopoietic replacement of beta cells. Science 311(5768):1778–1780 23. Yin D, Tao J, Lee DD, Shen J, Hara M, Lopez J, Kuznetsov A, Philipson LH, Chong AS (2006) Recovery of islet beta-cell function in streptozotocin-induced diabetic mice: an indirect role for the spleen. Diabetes 55(12):3256–3263

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24. Nishio J, Gaglia JL, Turvey SE, Campbell C, Benoist C, Mathis D (2006) Islet recovery and reversal of murine type 1 diabetes in the absence of any infused spleen cell contribution. Science 311(5768):1775–1778 25. Chong AS, Shen J, Tao J, Yin D, Kuznetsov A, Hara M, Philipson LH (2006) Reversal of diabetes in non-obese diabetic mice without spleen cell-derived beta cell regeneration. Science 311(5768):1774–1775 26. Melton DA (2006) Reversal of type 1 diabetes in mice. N Engl J Med 355(1):89–90 27. Lechner A, Yang YG, Blacken RA, Wang L, Nolan AL, Habener JF (2004) No evidence for signiﬁcant transdifferentiation of bone marrow into pancreatic beta-cells in vivo. Diabetes 53(3):616–623 28. Esni F, Stoffers DA, Takeuchi T, Leach SD (2004) Origin of exocrine pancreatic cells from nestin-positive precursors in developing mouse pancreas. Mech Dev 121(1):15–25 29. Delacour A, Nepote V, Trumpp A, Herrera PL (2004) Nestin expression in pancreatic exocrine cell lineages. Mech Dev 121(1):3–14 30. Dor Y, Brown J, Martinez OI, Melton DA (2004) Adult pancreatic beta-cells are formed by self-duplication rather than stem-cell differentiation. Nature 429(6987): 41–46 31. Georgia S, Bhushan A (2004) Beta cell replication is the primary mechanism for maintaining postnatal beta cell mass. J Clin Invest 114(7):963–968 32. Kushner JA, Ciemerych MA, Sicinska E, Wartschow LM, Teta M, Long SY, Sicinski P, White MF (2005) Cyclins D2 and D1 are essential for postnatal pancreatic beta-cell growth. Mol Cell Biol 25(9):3752–3762 33. Rane SG, Dubus P, Mettus RV, Galbreath EJ, Boden G, Reddy EP, Barbacid M (1999) Loss of Cdk4 expression causes insulin-deﬁcient diabetes and Cdk4 activation results in beta-islet cell hyperplasia. Nat Genet 22(1):44–52 34. Uchida T, Nakamura T, Hashimoto N, Matsuda T, Kotani K, Sakaue H, Kido Y, Hayashi Y, Nakayama KI, White MF, Kasuga M (2005) Deletion of Cdkn1b ameliorates hyperglycemia by maintaining compensatory hyperinsulinemia in diabetic mice. Nat Med 11(2):175–182 35. Gershengorn MC, Hardikar AA, Wei C, Geras-Raaka E, Marcus-Samuels B, Raaka BM (2004) Epithelial-to-mesenchymal transition generates proliferative human islet precursor cells. Science 306(5705):2261–2264 36. Ouziel-Yahalom L, Zalzman M, Anker-Kitai L, Knoller S, Bar Y, Glandt M, Herold K, Efrat S (2006) Expansion and redifferentiation of adult human pancreatic islet cells. Biochem Biophys Res Commun 341(2):291–298 37. Weinberg N, Ouziel-Yahalom L, Knoller S, Efrat S, Dor Y (2007) Lineage tracing evidence for in-vitro dedifferentiation, but rare proliferation, of mouse pancreatic β cells. Diabetes 56(5):1299–1304 38. Moss LG, Rhodes CJ (2007) Beta-cell regeneration: epithelial mesenchymal transition pre-EMTpted by lineage tracing? Diabetes 56(1):281–282 39. Chase LG, Ulloa-Montoya F, Kidder BL, Verfaillie CM (2007) Islet-derived ﬁbroblastlike cells are not derived via epithelial-mesenchymal transition from Pdx-1 or insulinpositive cells. Diabetes 56(1):3–7 40. Kulkarni RN, Jhala US, Winnay JN, Krajewski S, Montminy M, Kahn CR (2004) PDX1 haploinsufﬁciency limits the compensatory islet hyperplasia that occurs in response to insulin resistance. J Clin Invest 114(6):828–836 41. Finegood DT, Scaglia L, Bonner-Weir S (1995) Dynamics of beta-cell mass in the growing rat pancreas. Estimation with a simple mathematical model. Diabetes 44(3):249–256

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42. Scaglia L, Cahill CJ, Finegood DT, Bonner-Weir S (1997) Apoptosis participates in the remodeling of the endocrine pancreas in the neonatal rat. Endocrinology 138(4):1736–1741 43. Teta M, Long SY, Wartschow LM, Rankin MM, Kushner JA (2005) Very slow turnover of beta-cells in aged adult mice. Diabetes 54(9):2557–2567 44. Bouwens L, Lu WG, De Krijger R (1997) Proliferation and differentiation in the human fetal endocrine pancreas. Diabetologia 40(4):398–404 45. Sherr CJ, Roberts JM (2004) Living with or without cyclins and cyclin-dependent kinases. Genes Dev 18(22):2699–2711 46. Vermeulen K, Van Bockstaele DR, Berneman ZN (2003) The cell cycle: a review of regulation, deregulation and therapeutic targets in cancer. Cell Prolif 36(3): 131–149 47. Cozar-Castellano I, Weinstock M, Haught M, Velazquez-Garcia S, Sipula D, Stewart AF (2006) Evaluation of beta-cell replication in mice transgenic for hepatocyte growth factor and placental lactogen: comprehensive characterization of the G1/S regulatory proteins reveals unique involvement of p21cip. Diabetes 55(1):70–77 48. Cozar-Castellano I, Takane KK, Bottino R, Balamurugan AN, Stewart AF (2004) Induction of beta-cell proliferation and retinoblastoma protein phosphorylation in rat and human islets using adenovirus-mediated transfer of cyclin-dependent kinase4 and cyclin D1. Diabetes 53(1):149–159 49. Martin J, Hunt SL, Dubus P, Sotillo R, Nehme-Pelluard F, Magnuson MA, Parlow AF, Malumbres M, Ortega S, Barbacid M (2003) Genetic rescue of Cdk4 null mice restores pancreatic beta-cell proliferation but not homeostatic cell number. Oncogene 22(34):5261–5269 50. Fatrai S, Elghazi L, Balcazar N, Cras-Meneur C, Krits I, Kiyokawa H, Bernal-Mizrachi E (2006) Akt induces {beta}-cell proliferation by regulating cyclin D1, cyclin D2, and p21 levels and cyclin-dependent kinase-4 activity. Diabetes 55(2):318–325 51. Zhang X, Gaspard JP, Mizukami Y, Li J, Graeme-Cook F, Chung DC (2005) Overexpression of cyclin D1 in pancreatic beta-cells in vivo results in islet hyperplasia without hypoglycemia. Diabetes 54(3):712–719 52. Cozar-Castellano I, Fiaschi-Taesch N, Bigatel TA, Takane KK, Garcia-Ocana A, Vasavada R, Stewart AF (2006) Molecular control of cell cycle progression in the pancreatic beta-cell. Endocr Rev 27(4):356–370 53. Ohtsubo M, Theodoras AM, Schumacher J, Roberts JM, Pagano M (1995) Human cyclin E, a nuclear protein essential for the G1-to-S phase transition. Mol Cell Biol 15(5):2612–2624 54. Malek TR, Yu A, Scibelli P, Lichtenheld MG, Codias EK (2001) Broad programming by IL-2 receptor signaling for extended growth to multiple cytokines and functional maturation of antigen-activated T cells. J Immunol 166(3):1675–1683 55. Geng Y, Yu Q, Sicinska E, Das M, Schneider JE, Bhattacharya S, Rideout WM, Bronson RT, Gardner H, Sicinski P (2003) Cyclin E ablation in the mouse. Cell 114(4): 431–443 56. Bernal-Mizrachi E, Wen W, Stahlhut S, Welling CM, Permutt MA (2001) Islet beta cell expression of constitutively active Akt1/PKB alpha induces striking hypertrophy, hyperplasia, and hyperinsulinemia. J Clin Invest 108(11):1631–1638 57. Friedrichsen BN, Richter HE, Hansen JA, Rhodes CJ, Nielsen JH, Billestrup N, Moldrup A (2003) Signal transducer and activator of transcription 5 activation is sufﬁcient to drive transcriptional induction of cyclin D2 gene and proliferation of rat pancreatic beta-cells. Mol Endocrinol 17(5):945–958 58. Brockman JL, Schuler LA (2005) Prolactin signals via Stat5 and Oct-1 to the proximal cyclin D1 promoter. Mol Cell Endocrinol 239(1–2):45–53

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59. Kioussi C, Briata P, Baek SH, Rose DW, Hamblet NS, Herman T, Ohgi KA, Lin C, Gleiberman A, Wang J, Brault V, Ruiz-Lozano P, Nguyen HD, Kemler R, Glass CK, Wynshaw-Boris A, Rosenfeld MG (2002) Identiﬁcation of a Wnt/Dvl/beta-Catenin– >Pitx2 pathway mediating cell-type-speciﬁc proliferation during development. Cell 111(5):673–685 60. Chiles TC (2004) Regulation and function of cyclin D2 in B lymphocyte subsets. J Immunol 173(5):2901–2907 61. Bouchard C, Dittrich O, Kiermaier A, Dohmann K, Menkel A, Eilers M, Luscher B (2001) Regulation of cyclin D2 gene expression by the Myc/Max/Mad network: Mycdependent TRRAP recruitment and histone acetylation at the cyclin D2 promoter. Genes Dev 15(16):2042–2047 62. Krishnamurthy J, Ramsey MR, Ligon KL, Torrice C, Koh A, Bonner-Weir S, Sharpless NE (2006) p16INK4a induces an age-dependent decline in islet regenerative potential. Nature 443(7110):453–457 63. Karnik SK, Hughes CM, Gu X, Rozenblatt-Rosen O, McLean GW, Xiong Y, Meyerson M, Kim SK (2005) Menin regulates pancreatic islet growth by promoting histone methylation and expression of genes encoding p27Kip1 and p18INK4c. Proc Natl Acad Sci USA 102(41):14659–14664 64. Franklin DS, Godfrey VL, O’Brien DA, Deng C, Xiong Y (2000) Functional collaboration between different cyclin-dependent kinase inhibitors suppresses tumor growth with distinct tissue speciﬁcity. Mol Cell Biol 20(16):6147–6158 65. Pei XH, Bai F, Tsutsui T, Kiyokawa H, Xiong Y (2004) Genetic evidence for functional dependency of p18Ink4c on Cdk4. Mol Cell Biol 24(15):6653–6664 66. Latres E, Malumbres M, Sotillo R, Martin J, Ortega S, Martin-Caballero J, Flores JM, Cordon-Cardo C, Barbacid M (2000) Limited overlapping roles of P15(INK4b) and P18(INK4c) cell cycle inhibitors in proliferation and tumorigenesis. EMBO J 19(13):3496–3506 67. Moritani M, Yamasaki S, Kagami M, Suzuki T, Yamaoka T, Sano T, Hata J, Itakura M (2005) Hypoplasia of endocrine and exocrine pancreas in homozygous transgenic TGF-beta1. Mol Cell Endocrinol 229(1–2):175–184 68. Rachdi L, Balcazar N, Elghazi L, Barker DJ, Krits I, Kiyokawa H, Bernal-Mizrachi E (2006) Differential effects of p27 in regulation of beta-cell mass during development, neonatal period, and adult life. Diabetes 55(12):3520–3528 69. Fero ML, Rivkin M, Tasch M, Porter P, Carow CE, Firpo E, Polyak K, Tsai LH, Broudy V, Perlmutter RM, Kaushansky K, Roberts JM (1996) A syndrome of multiorgan hyperplasia with features of gigantism, tumorigenesis, and female sterility in p27(Kip1)-deﬁcient mice. Cell 85(5):733–744 70. Nakayama K, Ishida N, Shirane M, Inomata A, Inoue T, Shishido N, Horii I, Loh DY (1996) Mice lacking p27(Kip1) display increased body size, multiple organ hyperplasia, retinal dysplasia, and pituitary tumors. Cell 85(5):707–720 71. Georgia S, Bhushan A (2006) p27 Regulates the transition of beta-cells from quiescence to proliferation. Diabetes 55(11):2950–2956 72. Kaneto H, Kajimoto Y, Fujitani Y, Matsuoka T, Sakamoto K, Matsuhisa M, Yamasaki Y, Hori M (1999) Oxidative stress induces p21 expression in pancreatic islet cells: possible implication in beta-cell dysfunction. Diabetologia 42(9):1093–1097 73. Cozar-Castellano I, Haught M, Stewart AF (2006) The cell cycle inhibitory protein p21cip is not essential for maintaining beta-cell cycle arrest or beta-cell function in vivo. Diabetes 55(12):3271–3278 74. Kassem SA, Ariel I, Thornton PS, Hussain K, Smith V, Lindley KJ, Aynsley-Green A, Glaser B (2001) p57(KIP2) expression in normal islet cells and in hyperinsulinism of infancy. Diabetes 50(12):2763–2769

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75. Sempoux C, Guiot Y, Dahan K, Moulin P, Stevens M, Lambot V, de Lonlay P, Fournet JC, Junien C, Jaubert F, Nihoul-Fekete C, Saudubray JM, Rahier J (2003) The focal form of persistent hyperinsulinemic hypoglycemia of infancy: morphological and molecular studies show structural and functional differences with insulinoma. Diabetes 52(3):784–794 76. Zhang P, Liegeois NJ, Wong C, Finegold M, Hou H, Thompson JC, Silverman A, Harper JW, DePinho RA, Elledge SJ (1997) Altered cell differentiation and proliferation in mice lacking p57KIP2 indicates a role in Beckwith-Wiedemann syndrome. Nature 387(6629):151–158 77. Li FX, Zhu JW, Tessem JS, Beilke J, Varella-Garcia M, Jensen J, Hogan CJ, DeGregori J (2003) The development of diabetes in E2f1/E2f2 mutant mice reveals important roles for bone marrow-derived cells in preventing islet cell loss. Proc Natl Acad Sci USA 100(22):12935–12940 78. Iglesias A, Murga M, Laresgoiti U, Skoudy A, Bernales I, Fullaondo A, Moreno B, Lloreta J, Field SJ, Real FX, Zubiaga AM (2004) Diabetes and exocrine pancreatic insufﬁciency in E2F1/E2F2 double-mutant mice. J Clin Invest 113(10):1398–1407 79. Vasavada RC, Cozar-Castellano I, Sipula D, Stewart AF (2007) Tissue-speciﬁc deletion of the retinoblastoma protein in the pancreatic beta-cell has limited effects on beta-cell replication, mass, and function. Diabetes 56(1):57–64 80. Fajas L, Landsberg RL, Huss-Garcia Y, Sardet C, Lees JA, Auwerx J (2002) E2Fs regulate adipocyte differentiation. Dev Cell 3(1):39–49 81. Kulkarni RN, Bruning JC, Winnay JN, Postic C, Magnuson MA, Kahn CR (1999) Tissue-speciﬁc knockout of the insulin receptor in pancreatic beta cells creates an insulin secretory defect similar to that in type 2 diabetes. Cell 96(3):329– 339 82. Otani K, Kulkarni RN, Baldwin AC, Krutzfeldt J, Ueki K, Stoffel M, Kahn CR, Polonsky KS (2004) Reduced beta-cell mass and altered glucose sensing impair insulinsecretory function in betaIRKO mice. Am J Physiol Endocrinol Metab 286(1): E41–E49 83. Ueki K, Okada T, Hu J, Liew CW, Assmann A, Dahlgren GM, Peters JL, Shackman JG, Zhang M, Artner I, Satin LS, Stein R, Holzenberger M, Kennedy RT, Kahn CR, Kulkarni RN (2006) Total insulin and IGF-I resistance in pancreatic beta cells causes overt diabetes. Nat Genet 38(5):583–588 84. Kubota N, Terauchi Y, Tobe K, Yano W, Suzuki R, Ueki K, Takamoto I, Satoh H, Maki T, Kubota T, Moroi M, Okada-Iwabu M, Ezaki O, Nagai R, Ueta Y, Kadowaki T, Noda T (2004) Insulin receptor substrate 2 plays a crucial role in beta cells and the hypothalamus. J Clin Invest 114(7):917–927 85. Kubota N, Tobe K, Terauchi Y, Eto K, Yamauchi T, Suzuki R, Tsubamoto Y, Komeda K, Nakano R, Miki H, Satoh S, Sekihara H, Sciacchitano S, Lesniak M, Aizawa S, Nagai R, Kimura S, Akanuma Y, Taylor SI, Kadowaki T (2000) Disruption of insulin receptor substrate 2 causes type 2 diabetes because of liver insulin resistance and lack of compensatory beta-cell hyperplasia. Diabetes 49(11):1880–1889 86. Hashimoto N, Kido Y, Uchida T, Asahara S, Shigeyama Y, Matsuda T, Takeda A, Tsuchihashi D, Nishizawa A, Ogawa W, Fujimoto Y, Okamura H, Arden KC, Herrera PL, Noda T, Kasuga M (2006) Ablation of PDK1 in pancreatic beta cells induces diabetes as a result of loss of beta cell mass. Nat Genet 38(5):589–593 87. Stiles BL, Kuralwalla-Martinez C, Guo W, Gregorian C, Wang Y, Tian J, Magnuson MA, Wu H (2006) Selective deletion of Pten in pancreatic beta cells leads to increased islet mass and resistance to STZ-induced diabetes. Mol Cell Biol 26(7):2772–2781

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88. Tuttle RL, Gill NS, Pugh W, Lee JP, Koeberlein B, Furth EE, Polonsky KS, Naji A, Birnbaum MJ (2001) Regulation of pancreatic beta-cell growth and survival by the serine/threonine protein kinase Akt1/PKBalpha. Nat Med 7(10):1133–1137 89. Nakae J, Biggs WH, 3rd, Kitamura T, Cavenee WK, Wright CV, Arden KC, Accili D (2002) Regulation of insulin action and pancreatic beta-cell function by mutated alleles of the gene encoding forkhead transcription factor Foxo1. Nat Genet 32(2):245–253 90. Medema RH, Kops GJ, Bos JL, Burgering BM (2000) AFX-like Forkhead transcription factors mediate cell-cycle regulation by Ras and PKB through p27kip1. Nature 404(6779):782–787 91. Schmidt M, Fernandez de Mattos S, van der Horst A, Klompmaker R, Kops GJ, Lam EW, Burgering BM, Medema RH (2002) Cell cycle inhibition by FoxO forkhead transcription factors involves downregulation of cyclin D. Mol Cell Biol 22(22): 7842–7852 92. Kitamura T, Nakae J, Kitamura Y, Kido Y, Biggs WH, 3rd, Wright CV, White MF, Arden KC, Accili D (2002) The forkhead transcription factor Foxo1 links insulin signaling to Pdx1 regulation of pancreatic beta cell growth. J Clin Invest 110(12): 1839–1847 93. Kushner JA, Ye J, Schubert M, Burks DJ, Dow MA, Flint CL, Dutta S, Wright CV, Montminy MR, White MF (2002) Pdx1 restores beta cell function in Irs2 knockout mice. J Clin Invest 109(9):1193–1201 94. Biondi CA, Gartside MG, Waring P, Lofﬂer KA, Stark MS, Magnuson MA, Kay GF, Hayward NK (2004) Conditional inactivation of the MEN1 gene leads to pancreatic and pituitary tumorigenesis but does not affect normal development of these tissues. Mol Cell Biol 24(8):3125–3131 95. Bertolino P, Tong WM, Herrera PL, Casse H, Zhang CX, Wang ZQ (2003) Pancreatic beta-cell-speciﬁc ablation of the multiple endocrine neoplasia type 1 (MEN1) gene causes full penetrance of insulinoma development in mice. Cancer Res 63(16): 4836–4841 96. Pelengaris S, Khan M, Evan GI (2002) Suppression of Myc-induced apoptosis in beta cells exposes multiple oncogenic properties of Myc and triggers carcinogenic progression. Cell 109(3):321–334 97. Lawlor ER, Soucek L, Brown-Swigart L, Schhors K, Bialucha CU, Evan GI (2006) Reversible kinetic analysis of Myc targets in vivo provides novel insights into Myc-mediated tumorigenesis. Cancer Res 66(9):4591–4601 98. Vasavada RC, Gonzalez-Pertusa JA, Fujinaka Y, Fiaschi-Taesch N, Cozar-Castellano I, Garcia-Ocana A (2006) Growth factors and beta cell replication. Int J Biochem Cell Biol 38(5–6):931–950 99. Heit JJ, Karnik SK, Kim SK (2006) Intrinsic regulators of pancreatic beta-cell proliferation. Annu Rev Cell Dev Biol 22:311–338 100. Heit JJ, Apelqvist AA, Gu X, Winslow MM, Neilson JR, Crabtree GR, Kim SK (2006) Calcineurin/NFAT singalling regulates pancreatic beta-cell growth and function. Nature 443:345–349

14. Stem Cells as a Cure for Diabetes Timo Otonkoski, Meenal Banerjee, and Karolina Lundin

Summary. Clinical islet transplantation trials have shown that it is possible to at least partially restore functional beta-cell mass in type 1 diabetic patients. However, the scarcity of organ donors and need for intensive immunosuppression prevent the widespread use of this treatment regimen. Progress in stem cell biology provides hope for the solution of these problems. New beta cells could be grown in the laboratory from pancreatic precursors or from more multipotent stem cells. Neogenesis of islets from intrapancreatic precursors has been demonstrated in a number of studies. Fully differentiated beta cells have only been derived in primary cultures, mainly reﬂecting transdifferentiation from acinar or ductal cells. It is unlikely that these cells could provide a quantitatively sufﬁcient source for cell transplantation. However, the stimulation of endogenous beta-cell regeneration is a promising therapeutic possibility. Extensive proliferation of supposed pancreatic tissue stem cells has also been found, but full maturation of the expanded cells into functional islet cells has not been reported. The most promising possibility to generate theoretically unlimited numbers of beta cells is based on the use of embryonic stem (ES) cells. Human ES cells can be taken through a multiphase differentiation protocol to become insulin-expressing cells that phenotypically resemble immature human fetal beta cells. This ﬁeld is still in its infancy and many problems need to be solved before ES-cell based products could be developed to the level required for clinical trials in diabetes.

Background Type 1 diabetes results from the absolute loss of pancreatic beta cells. Consequently, the optimal treatment for this condition would be to restore the functional beta-cell mass. So far, the only clinically applicable means towards Hospital for Children and Adolescents and the Biomedicum Stem Cell Center, Biomedicum Helsinki, P.O. Box 63, Haartmaninkatu 8, FI-00014 University of Helsinki, Helsinki, Finland

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this goal has been the transplantation of the pancreas or islets puriﬁed from organ donor pancreata, an approach that can never be applied to a large number of patients. Recent results in experimental animal models and cultured human pancreatic cells suggest that the regenerative capacity of the pancreatic islets may be much greater than previously anticipated. Research efforts are aimed at the identiﬁcation and isolation of the islet precursor cells and molecular mechanisms of their expansion and differentiation. At the same time, research on pluripotent human embryonic stem cells has emerged and is growing exponentially. There is reason for optimism also for these stem cells as a potentially unlimited source of physiologically functioning human insulin-producing cells that could in the future be applied therapeutically. In the following chapter, the current status of these various research ﬁelds is brieﬂy reviewed.

Transplantation of Beta Cells: Where Are We? Restoration of insulin secretion through transplantation of insulinproducing cells in a diabetic patient has long been a dream, supported by the excellent efﬁcacy of islet transplantation in syngeneic animal models. The results of early clinical trials were disappointing, where insulin independence was obtained in less than 10% of islet recipients [1]. The so-called Edmonton protocol, involving the intraportal infusion of islets from two or more organ donors under glucocorticoid-free immunosuppression, showed that independence of injected insulin could be obtained in seven consecutive patients [2]. Further studies and longer follow-up times have shown that insulin independence is gradually lost in most cases, but even partial graft function provides signiﬁcant beneﬁt to the patients [3]. Currently, islet transplantation is considered as an experimental therapy, only applicable in clinical trials in highly selected patients. Nevertheless, these studies have provided the proof of principle for beta-cell replacement; it is possible to restore long-term beta-cell function by cell transplantation in patients with type 1 diabetes and to a degree providing better glycemic control than any conventional treatment. However, it is obvious that the number of islets needed far exceeds the source of islets from deceased donors. Also the side effects of the immunosuppressive therapy are unacceptable for widespread application of this therapy in type 1 diabetes. Alternative sources of cells are needed and the capacity of islet regeneration from endogenous precursor cells should be exploited together with approaches to halt beta-cell targeted autoimmunity.

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Islet Precursors in the Pancreas: What Are They and Could They be Employed Therapeutically? The mass of beta cells changes in response to the insulin demand of the mammalian organism (see the chapter by Rhodes in this volume). Assuming that autoimmunity could be controlled in type 1 diabetes, beta cells might regenerate to a much higher degree than previously anticipated, as suggested in the NOD mouse model of autoimmune diabetes [4–7]. Identiﬁcation of the molecular mechanisms controlling the regenerative process, and the exact identity of the cells that can serve as islet precursors remain the most important aims of current research efforts. Beta-cell proliferation is apparently the most important mechanism for the increase of beta-cell mass in adult rodents [8]. The capacity of human beta cells to proliferate is much lower [9,10]. Several experimental models of pancreatic regeneration have provided indirect evidence for the neogenesis of islets that appear to bud out of the ductal epithelium [11–14]. Morphological evidence suggestive of islet neogenesis can often be detected in sections of the human, particularly diabetic, pancreas [15–17]. Despite a lot of research, the existence and exact nature of the islet precursor cell remains elusive and controversial. Studies aiming to identify and characterize the pancreatic islet precursor cells are summarized in Table 1. These can be roughly divided in two categories. The ﬁrst of these deals with plasticity of pancreatic cell types, often describing differentiation of islet cells from cytokeratin-expressing ductal epithelial cells or transdifferentiation of acinar cells into ductal and islet cells [18–29]. These processes are associated with no or only limited expansion of the cell mass. The second category consists of studies demonstrating extensive growth of putative islet precursor cells [30–36]. In contrast to the shortterm cultures, the endocrine differentiation of the extensively expanded precursor cells is often limited. As can be seen in Table 1, there is remarkable variability in the characteristics of the islet precursor cells among various experiments. Ramiya and colleagues prepared long-term cultures of pancreatic duct cells from adult prediabetic NOD mice and could show that these cells were able to differentiate into all major islet cell types and reverse diabetes after implantation in to diabetic NOD mice [30]. At the same time, Bonner-Weir and colleagues reported in vitro generation of islets from human ductal cells after limited expansion [18]. The same model was used by Gao and colleagues who showed that islet neogenesis was strongly inhibited by serum and stimulated by nicotinamide and extracellular matrix [20]. The same group later

Isolated islets

Islet depleted, duct rich fraction

Isolated islets

Islets and duct cells

Canine and human

Human

Mouse

Mouse

Petropavlovskaia and Rosenberg, 2002 [19]

Gao et al., 2003 [20]; 2005 [21]; 2006 [22]

Banerjee and Bhonde, 2003 [23]

Seaberg et al., 2004 [24]

A. Studies showing no or only limited cell expansion Human Bonner-Weir et al., Islet depleted, 2000 [18] duct rich fraction CK 19 positive ductal epithelium with cytoplasmic or weak nuclear expression of Ipf1 Small cells positive for Pdx1, synaptophysin and hormones but negative for CK19 and nestin CK 19 and NCAM positive ductal epithelium with cytoplasmic expression of Ipf1 IIPCs (Intra Islet Precursor Cells) positive for CK7 and CK19 but negative for insulin PMPs (Pancreasderived Multipotent Precursors) express both pancreatic and neural precursor markers

Not performed

Not performed

Limited expansion

Morphological maturation after implantation in mice

Not performed

Not performed

Cell expansion for single generation

Cell expansion for single generation

Limited and very low rate of proliferation

Cell expansion for single generation

Table 1. Reports published since the year 2000 focusing on in vitro characterization, expansion and differentiation of pancreas-derived islet precursor cells Authors, year [Ref.] Characteristics of the Expansion In vivo studies Species Source of cells islet precursor

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Pancreatic exocrine acini

Pancreatic exocrine cells

Islet-depleted mixed pancreatic epithelial cells

Non-islet fraction consisting of less than 1% beta cells Embryonic pancreas (E15.5)

Rat

Mouse

Human

Human

Mouse

Baeyens et al., 2005 [25]

Minami et al., 2005 [26]

Hao et al., 2006 [27]

Todorov et al., 2006 [28]

Sugiyama et al., 2007 [29]

NGN3+ islet progenitor cells expressing CD133 and CD49f

Acinar cells lost expression of amylase and started to express CK20 in suspension culture Genetic lineage tracing shows that insulinsecreting cells were derived from acinar cells NEPEC (Non Endocrine Pancreatic Epithelial Cells), CK 19 positive, coexpression of vimentin and nestin in monolayers Cells upon expansion expressed CK7/19 with nuclear and cytoplasmic expression of Ipf1 Enriched and cultured, not expanded in vitro

Cells were expanded to form conﬂuent monolayer

Limited expansion

Limited expansion

Acinar cells were expanded to form a monolayer

In vitro differentiated ICCs further mature upon transplantation into mice Not performed

Beta-cell differentiation only when cotransplanted in mice with fetal pancreas

Reversal of diabetes in mice by transplantation of transdifferentiated cells Not performed

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Species Source of cells

Isolated islets

Single cell suspension of neonatal and adult pancreas

Human

Mouse

Gershengorn et al., 2004 [32]

Suzuki et al., 2004 [33]

Isolated islets

Rat and human

Zulewski et al., 2001 [31]

B. Studies showing extensive precursor cell expansion Mouse Ducts from preRamiya et al., 2000 diabetic NOD [30] mice

Authors, year [Ref.]

Table 1. Continued

hIPCs (human Isletderived Precursor Cells) positive for vimentin, nestin and SMA PSCs (Pancreatic Stem Cells) express C-Met but are negative for hematopoietic and endothelial antigens

IPSC/IPC (Islet Producing Stem Cell/Islet Progenitor Cell) positive for many islet markers NIP (Nestin-positive Islet-derived Progenitor) cells negative for CK19

Characteristics of the islet precursor

Extended expansion with self renewing cell divisions in culture

Extended proliferation, maintained approximately for 8 months Extensive expansion after epithelial-tomesenchymal transition

Long-term culture (more than 3 years)

Expansion

Redifferentiated cells released Cpeptide upon implantation in mice Transplanted cells differentiated into pancreatic endocrine and acinar cells

Reduction of plasma glucose upon implantation of in vitro generated islets in NOD mice Not performed

In vivo studies

270 T. Otonkoski et al.

Human

Mouse

Ouziel-Yahalom et al., 2006 [35]

Yamamoto et al., 2006 [36] Pancreatic ductal cells

Dispersed pancreatic islets

Pancreatic duct tissue

Putative PSCs (Pancreatic Stem Cells) expressing Pdx1, nestin and mesenchymal stem cell markers PHID (Proliferating Human Islet Derived) cells expressed vimentin PPPD (Pdx-1 Positive Pancreatic Ductderived) cells expressing high level of CK19, CAII and Pdx1 Expansion over 65 000 fold (16 population doublings) Maintenance of cells in repeated passages for more than a year in presence of cholera toxin

Expanded for more than 20 cell passages

Not performed

Not performed

Not performed

CK19, cytokeratin 19; CAII, carbonic anhydrase II; Ipf1, insulin promoter factor 1 (human); Pdx1, pancreatic duodenal homeobox gene 1 (mouse); NGN3, neurogenin 3; SMA, smooth muscle actin

Human

Lin et al., 2006 [34]

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showed that islet cell differentiation in this culture model appeared to depend on N-CAM positive cells many of which co-expressed cytokeratin 19 [21]. In 2001, Zulewski and coworkers described long-term culture of isletderived progenitor cells cultured from rat and human islets [31]. These cells were characterized by their expression of nestin, an intermediate ﬁlament protein that had been mainly associated with neuronal precursors. Other studies have also identiﬁed nestin expression in the proliferating islet precursor cells [32,34]. Analysis of embryonic pancreas development and cell lineage tracing experiments have clearly shown that in vivo, islet cell precursors never express nestin [37–40]. The proliferating nestin-positive islet precursor cells which have mixed epithelial-mesenchymal phenotypes may thus represent a phenomenon speciﬁc for the culture conditions. The true precursor capacity of these cells remains uncertain because only a very limited level of endocrine re-differentiation has been demonstrated and there is no evidence of differentiation into insulin-producing cells of potential therapeutic value [32,34,35].

Bone Marrow-Derived Stem Cells Several nonpancreatic cell sources have also been proposed to be able to differentiate into islet cells. The potential of hepatocytes is discussed elsewhere (see chapter by Ferber in this volume). In addition to hematopoietic stem cells, bone marrow (BM) contains stromal or mesenchymal stem cells which can readily be differentiated in vitro into a variety of mesenchymal cell types providing obvious therapeutic possibilities [41]. Although there have been some reports clearly suggesting that BM-derived stem cells could directly differentiate into islet cells in vivo [42] or in vitro [43], there is now general consensus that this does not occur to a reproducibly measurable degree [44–46]. Instead, there is an increasing number of reports suggesting that BM-derived stem cells, either mesenchymal or hematopoietic or even transplanted splenocytes, may have a potent stimulatory effect on pancreatic islet regeneration [4–6,44,45,47,48]. An intriguing possibility is that the spleen may contain a population of multi-lineage stem cells that are not found in the bone marrow but are able to differentiate into pancreatic islet cells [7,49,50]. Stimulation of endogenous beta-cell regeneration through the homing of transplanted stem cells into the pancreas is a rapidly evolving ﬁeld of investigation carrying obvious possibilities for clinical application in conjunction with measures to control beta-cell targeted autoimmunity.

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Embryonic Stem Cells: First Steps Taken on a Long and Winding Road When the ﬁrst human embryonic stem (ES) cell lines were derived in 1998 [51], they were hoped to provide a solution for the shortage of cells for cell replacement therapies for a number of degenerative diseases, including diabetes. By that time, almost 20 years of research on mouse ES cells had already shown that ES cells have the potential to differentiate into physiologically functioning cells of various types. However, there are some critical differences between human and murine ES cells, even involving different molecular mechanisms to maintain the state of pluripotency [52]. No direct extrapolations can thus be made. Embryonic stem cells are derived from the inner cell mass (ICM) of 5–6day-old mammalian blastocysts, in the case of human, spare embryos from in vitro fertilization. The ICM contains the cells that in a normally developing embryo would form the embryo proper. Hence, these cells retain the property to form all cells of an organism. Under proper conditions, the ES cells can be kept in culture virtually indeﬁnitely without losing their pluripotency, thus creating a theoretically endless source of non-transformed cells with the potential to be differentiated into the desired direction [53]. Both mouse and human ES cells can give rise to insulin producing cells when allowed to form three-dimensional structures called embryoid bodies (EBs) in suspension culture. EBs are considered to recapitulate the mechanisms occurring during gastrulation and contain cells of all three germ layers, including endoderm [54]. A low number of insulin-positive cells can be detected within the EBs [55–59]. Using cell-trap systems with drug resistance genes under the insulin or Nkx6.1 promoters, enough insulin-positive cells could be isolated, expanded and transplanted in diabetic mice, resulting in normalization of blood glucose levels, indicating the functionality of these mESC-derived cells [55,58]. Several studies have aimed at increasing the number of mouse ES cellderived insulin-producing cells and their functional maturity either by modifying culture conditions, by genetic manipulations, or a combination of both. In 2001, a modiﬁcation of a protocol used for neuronal differentiation was reported to result in efﬁcient generation of pancreatic islet-like cells [60]. This protocol was further modiﬁed in a number of ways, including addition of phosphatidylinositol kinase (PI3K) inhibitors [61], overexpression of Pax4 or Pdx1 [62,63], or minor modiﬁcations of the culture conditions [64,65]. These results have been controversial, and sharp criticism has been expressed against the “neuroectoderm-based” protocols. It has been shown that insulinpositive cells found in these cultures may represent apoptotic cells passively

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taking up insulin from the medium which contains a high concentration of exogenous insulin [66]. Criticism has also been targeted against the principle of forming beta cells via ectoderm, as opposed to the embryologically correct deﬁnitive endoderm [67,68]. The necessity of the EB formation step has been debated. Some investigators have shown that endoderm is only generated via an EB stage whereas monolayer cultures select for ectoderm [56,69–71], while others claim that the endodermal derivatives formed via the EB stage are of visceral, not deﬁnitive endoderm origin [72,73]. It has, however, also been shown that deﬁnitive endoderm commitment occurs within the EBs [74–76]. At this stage it is important to stress the need for thorough phenotypic analysis of ES derived cells. In the case of insulin-producing cells the analysis must include not only quantitative analysis of insulin gene and protein expression, but also evidence of its proper processing, expression of key proteins needed for glucose sensing and regulated exocytosis, as well as analyses of insulin release induced by glucose and other secretagogues. None of the studies reported until now on the derivation of insulin-secreting cells from mouse ES cells has properly demonstrated the essential characteristics required for a normal beta cell. It is obvious that any stem-cell derived cell intended to be used as a physiological insulin delivery vehicle needs to fulﬁll at least the essential metabolic and electrophysiologic features of a beta cell [77]. It seems inevitable that in order to achieve this, the major developmental steps involved in the differentiation pathway to a beta cell in vivo have to be mimicked. One of the major problems has been the lack of markers that can distinguish deﬁnitive from extraembryonic endoderm. The ﬁrst undisputable evidence that deﬁnitive endoderm (DE) can be generated from mouse ES cells came in 2004, when Kubo and co-workers used cells expressing GFP under the control of the brachyury promoter, a transcription factor expressed in the primitive streak and later in mesoderm. They showed that endodermal cells were generated from ES cell-derived primitive streak mesoderm cells, and hence, could not be of extraembryonic origin [74]. Low serum concentration and addition of activin were crucial for the formation of DE. It is now generally agreed that no single marker can reliably deﬁne DE cells. Instead, an entire panel of transcription factors and cell surface proteins needs to be analysed [74,76,78–80]. Some of the markers that can be used for this purpose are listed in Table 2. Later studies have shown that in addition to activin, retinoic acid is required for commitment of the pancreatic endoderm [75,81,82]. In 2005, Nishikawa and collaborators published a study on mouse ES cells in which they, in addition to describing a method of generating a pure population of DE cells from mouse ES cells after Activin A induction, also identi-

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Table 2. Some markers used to deﬁne deﬁnitive endoderm lineage in differentiating ES cells Gene Expression in Refs. Foxa2 (hepatocyte nuclear factor 3beta) Sox17 (sex determining region Y-box 17) Cer (Cerberus 1) Cxcr4 (chemokine C-X-C motif receptor 4) GSC (goosecoid) Sox7 (sex determining region Y-box 7) AFP (alpha-fetoprotein) NCAD (N-cadherin)

DE, EE, PS and mesoderm DE and EE DE, EE, mesoderm DE and mesoderm, not in EE DE, EE, PS and mesoderm EE, not in DE EE, not in early DE but later in hepatocytes DE, PS, mesoderm, not in EE

[95–97] [98] [99–102] [103] [104] [98] [105,106] [107–109]

DE, deﬁnitive endoderm; EE, extraembryonic endoderm (including primitive-, visceral-, and parietal endoderm); PS, primitive streak

ﬁed CXCR4 as a cell surface marker expressed in DE at this stage. This cytokine receptor was the ﬁrst identiﬁed surface marker speciﬁc for DE, and provided a tool not only to more easily identify, but also to isolate DE cells [76,78]. This paper was soon followed by a study by D’Amour and collaborators on human ES cells also using restricted serum and high dose activin for derivation of DE [80]. It was later shown that inhibition of the insulin/IGF-1 signalling pathway by addition of a PI3K inhibitor could effectively substitute for the reduced serum concentration in their protocol [83]. The problem seemed to be the reluctance of the DE cells derived from both mouse and human ES cells to differentiate further, and speciﬁcally into a pancreatic fate. In particular, FACS-sorted, isolated DE cells were not able to differentiate further, highlighting the importance of intercellular interactions in a heterogeneous differentiating culture. Several reports have shown that either coculture or cotransplantation of partially differentiated ES cells with embryonic pancreatic rudiments from mouse drastically increases the differentiation of pancreatic endocrine cells [58,79,84,85]. The same is true also for the induction of adult human pancreatic duct cell differentiation into islets [27]. Recently D’Amour and coworkers described a reﬁned and extended protocol resulting in differentiation of human ES cells into insulin producing cells [86]. For the ﬁrst time, it appears that true immature beta cells had been derived from human ES cells. In this 5-stage, 16-day differentiation protocol, knowledge from in vivo differentiation of pancreatic beta cells was utilized, and human ES cells were taken through a number of consecutive steps mimicking those occurring physiologically (Fig. 1). In the ﬁrst stage, DE was induced by a very low serum concentration in addition to a high concentration of Activin A and Wnt3a. In the second stage, a primitive gut tube was

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Fig. 1. Schematic presentation of a ﬁve-stage protocol for the differentiation of pancreatic endocrine cells from hESCs through intermediate endodermal stages. The culture media and growth factors used at each stage are shown together with some marker characteristics of each cell population. CYC, KAAD-cyclopamine; RA, all-trans-retinoic acid; DAPT, gamma-secretase inhibitor; Ex4, exendin-4; ME, mesendoderm; DE, deﬁnitive endoderm; PG, primitive gut tube; PF, posterior foregut endoderm; PE, pancreatic endoderm and endocrine precursor; EN, endocrine cell. Reprinted from D’Amour and colleagues [86] with permission from Macmillan Publishers Ltd

induced by withdrawal of activin and addition of the sonic hedgehog inhibitor KAAD-cyclopamine (Cyc) and ﬁbroblast growth factor 10 (FGF10). Following this, posterior foregut was induced by retinoic acid, and pancreatic endoderm and endocrine precursor cell formation by the glucagon-like peptide-1 (GLP1) analog exendin-4 and a Notch signalling (gamma-secretase) inhibitor. Further maturation of islet cells was stimulated by addition of exendin-4, insulin-like growth factor 1 (IGF1) and hepatocyte growth factor (HGF). Although the protocol described above is undoubtedly a major breakthrough for the induction of beta cells from human ES cells, it has to be underlined that mature beta cells have not yet been achieved. For example, insulin release from these immature beta cells was not glucose responsive and it has not yet been shown that the cells can rescue diabetic mice. Also, important quantitative problems need to be solved because the endocrine precursor cells do not proliferate and maximally only 7% of the differentiated cells gained a beta-cell phenotype. Hence, very large numbers of human ES cells would be needed to produce sufﬁcient quantities of insulin-secreting cells for transplantation in large animals or humans. Based on the recent remarkable progress, it now seems likely that a source of human ES cell-derived physiological insulin-producing cells will be developed. Even after that, many issues remain to be solved before clinical trials could be planned. One of them concerns the tumorigenicity of the ES cells. Upon transplantation into immunocompromised mice, ES cells form

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teratomas, a phenomenon used as proof for their pluripotency. It has been shown that ES cells that are differentiated in vitro lose their ability to form teratomas [87]. However, ES-derived cells are likely to be heterogeneous, and may contain stem cells that have escaped differentiation. Robust methods are needed to ensure that such cells are completely eliminated from preparations intended for therapeutic use. Another safety concern is the contamination of human ES cell lines by xenogenic antigens from poorly characterized components used for their culture, such as feeder cells, extracellular matrix preparations and serum replacement solutions [88,89]. To make them useful for transplantation purposes, cell lines derived and maintained according to good manufacturing practice (GMP) standards are required. Cell lines meeting these criteria are now being developed [90]. Several hundred human ES cell lines currently exist worldwide. Although all the lines share major characteristics, there are some basic differences between the lines [91], most importantly in their spontaneous differentiation patterns [92]. This could be important for their suitability for certain differentiation protocols and function upon transplantation. Finally, an important issue remains concerning the immunogenicity and recurrence of autoimmunity against stem cell-derived beta cells after transplantation. Until now, it has been impossible to address these issues in human type 1 diabetes. It is known that both allograft rejection and re-activation of beta-cell autoimmunity are important reasons for the loss of islet graft function [1,93]. While there are some indications from experimental models that ES-cell derived grafts could have immunological beneﬁts [94], it has so far not been possible to properly study this using ES-derived beta cells. Theoretically, a stem cell-based source provides obvious beneﬁts as compared with the transplantation of primary beta cells, because of the possibility to genetically modify the stem cells to reduce their immunogenicity. Acknowledgments. The authors are grateful to Dr. Päivi Miettinen for valuable comments on the manuscript. Our studies related to this review have been supported by the Juvenile Diabetes Research Foundation, the Academy of Finland, and the Sigrid Jusélius Foundation.

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15. Use of Extra-Pancreatic Tissues for Cell Replacement Therapy for Diabetes Irit Meivar-Levy* and Sarah Ferber†,*

Summary. Pancreatic islet transplantation may constitute the best approach for the long-term control of blood glucose levels in the treatment of diabetes. However, tissue replacement therapy will become widely available as a treatment for diabetic patients only when islets or insulin-producing cells are available in unlimited amounts, and will not be rejected by the diabetic recipients. The present chapter will analyze the option of using adult extrapancreatic tissues for regulated insulin production. Two major approaches could endow adult extra-pancreatic tissues with characteristics and functions that can be used for diabetes cell replacement therapy: First, insulin gene therapy, which involves the ectopic expression of constructs encoding the proinsulin gene or modiﬁed proinsulin sequences in adult extra-pancreatic cells from different sources. Second, the induction of a process called developmental redirection of extra-pancreatic tissues into insulin-producing cells. The second approach exploits the instructive roles of pancreatic transcription and soluble factors in controlling pancreas organogenesis in the embryo to dictate the induction of pancreatic lineage and function also in adult tissues. This approach endows adult extra-pancreatic tissues with pancreatic characteristics and function, thus promoting ex vivo differentiation into insulinproducing and secreting cells. Using adult extra-pancreatic tissues may result in the generation of custom made “self” surrogate pancreatic beta cells for the treatment of diabetes, bypassing both the shortage in tissue availability from cadaveric donors and the need for life-long immunosuppression. *Endocrine Institute, Sheba Medical Center, Tel-Hashomer 52621, Israel † Department of Human Genetics and Molecular Medicine, Sackler School of Medicine, Tel-Aviv University, Tel-Aviv, Israel

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Introduction Pancreas transplantation and islet-cell implantation are effective in replacing the function of the impaired tissue and inducing continuous normoglycemia. These grafts bring freedom from the burden of insulin injections, glucose testing and dietary restrictions; and, importantly, continuous normoglycemia protects against the complications of diabetes [1]. However, these treatments require extensive, lifelong suppression of the immune system and are restricted by the limited tissue supply from cadaveric donors. Generally, it is accepted that islet cell implantation will become widely available only when new sources of islets or pancreatic beta cells are found [2–5]. Since differentiated beta cells cannot be efﬁciently expanded in vitro [6], alternative approaches are being explored to generate insulin-producing cells. Embryonic stem cells [7–10] have the theoretical potential to serve as an expandable pool of cells that can be induced toward insulin production and beta cell phenotype with the right culture conditions. The ethical controversies and the oncogenic potential are still major obstacles to be overcome before insulin-producing cells derived from embryonic sources could be used for therapy [11,12]. The present chapter will analyze the option of using adult extra-pancreatic tissues for insulin production. There are two major approaches by which adult extra-pancreatic tissues can be used for diabetes cell replacement therapy: 1. Insulin gene therapy, which involves the ectopic expression of constructs encoding the proinsulin gene or modiﬁed proinsulin sequences in adult extra-pancreatic cells from different sources [13,14]. 2. The induction of a process called developmental redirection of extrapancreatic tissues into insulin-producing cells. This approach uses the instructive roles of pancreatic transcription factors in controlling pancreas organogenesis in the embryo and exploits them to dictate the induction of pancreatic lineage and function also in adult tissues. Indeed, the ectopic expression of pancreatic transcription and growth factors has been implicated in inducing the expression of otherwise silent pancreatic genetic information in adult, extra-pancreatic tissues [15]. This process endowed the adult extra-pancreatic tissues with pancreatic characteristics and function. While in the ﬁrst approach the ectopic gene expression endows the extrapancreatic cells with narrow beta-cell like characteristics, mainly manifested by the capacity to produce insulin, wide alterations in the host repertoire of gene expression are induced in the second approach. The concerted activation of numerous otherwise silent pancreatic genes allows a good simulation of normal beta-cell function, upon developmental redirection.

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Using adult extra-pancreatic tissues may result in the generation of custom made “self” surrogate beta cells for the treatment of diabetes, bypassing both the shortage in tissue availability from cadaveric donors and the need for lifelong immunosuppression.

Insulin Gene Therapy Potential Adult Tissues Sources and Gene Delivery Tools The beta cell clearly is one of, if not the most intricate cellular machines having to cope with a variety of conditions where physiological adaptation is required. Beta cells are able to synthesize, process, package, store and rapidly turn on and off vesicular insulin secretion in tight adherence to glucose levels [16,17]. It is clear that simple replacement of insulin gene expression by genetic engineering is not likely to be useful as a therapy for type 1 diabetes unless it includes an appropriate system to regulate insulin secretion in response to blood glucose levels. Several cells sources have been suggested as possible targets for insulin production and secretion, due to their characteristics that allow a partial simulation of beta-cell function (as summarized in Table 1): 1. Neuroendocrine cells, like pituitary cells, intestinal K and L cells and gastric G cells, since these cells have a regulated secretory pathway for protein secretion and enzymatic machinery for processing precursor proteins into active molecules [18,19]. 2. Muscle cells, since these cells play a key role in glucose homeostasis, and express the endopeptidase furin which is capable of protein processing [20–22]. 3. Hepatocytes, because they have glucose-sensing molecules similar to those in pancreatic beta cells, such as Glut2 and glucokinase. In addition, a number of glucose regulatable hepatocyte gene promoters are available to drive the insulin gene expression [23,24]. Due to the chronic nature of disease, gene therapy for type 1 diabetes requires long-term transgene expression [25]. Currently, only a few vectors are able to mediate long-term transgene expression in post-mitotic cells with reasonable safety proﬁles [26–28]. The most commonly used vehicles for genes delivery into mammalian cells exploit the high infectivity of DNA or RNA based viruses that have been engineered to express the gene of interest and lack replication capacity: • Retroviral vectors are derived from RNA viruses: These viruses main feature is the reverse transcriptase activity which enables them to transcribe their

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Table 1. Tissues for insulin gene therapy Cell/tissue Major advantages

Main disadvantages

References

Pituitary cells

Regulated secretory pathway Enzymatic machinery for processing precursor proteins

No glucose sensing abilities Co-secretion of ACTH

[39–43]

Intestinal derived cells (K & L cells)

Regulated secretory pathway Glucose sensing abilities Express PC1/3 and PC2

Rare population Low accessibility

[45–48]

Gastric derived G cells

Regulated secretory pathway Glucose sensing abilities Express PC1/3 and PC2

Rare population Low accessibility

[52]

Muscle cells

High accessibility Capacity to produce large amounts of secreted proteins Key role in glucose homeostasis

No glucose sensing abilities No regulated secretory pathway No enzymatic machinery for processing pro-insulin

[57–67,69]

Liver cells

Moderate accessibility Key role in glucose homeostasis Glucose sensing abilities Glucose regulate gene promoters

No enzymatic machinery for processing pro-insulin No rapid response to changes in blood glucose levels

[71–76, 80–84, 86–88]

viral RNA genome into a double-stranded viral DNA, which then is stably inserted into the host genome [29]. These vectors are widely used systems for the introduction of foreign genes into target cells. About 40% of the clinical gene therapy protocols are performed using recombinant retroviruses. However, these vectors are inefﬁcient in expressing genes in nondividing cells. • Lentiviral vectors: The human immunodeﬁciency virus (HIV) is a lentivirus and causes acquired immune deﬁciency syndrome (AIDS). This type of retrovirus is of interest since it has the capacity to infect and integrate into nondividing cells [30,31]. This ability of lentiviruses is due to their capacity

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to enter the nucleus without the requirement of mitosis. However, lentiviruses achieve stable transduction by integrating randomly into the host genome, which raises the possibility of insertional mutagenesis following vector delivery. • Recombinant adeno-associated virus (r-AAV) vectors: AAV is a family of single-stranded DNA viruses that belong to the family of Parvoviruses. Broad ranges of cell types are susceptible to AAV infection and the recombinant AAVs are nonimmunogenic vectors, able to stably transduce both dividing and nondividing cells. However, due to their relatively small packaging size, rAAV has a limited capacity (<4.8 kb) for transgene insertion [32,33]. Theoretically AAV integrate in a speciﬁc site on the host genome located in chromosome 19. • Recombinant adenoviruses are nonenveloped DNA viruses carrying linear double stranded DNA of about 35 kb: About 50 distinct serotypes of distinguishable viruses are known, but the most commonly used in gene therapy are the serotypes 2 and 5. Recombinant adenoviruses efﬁciently infect nondividing cells. However, since these viruses do not integrate into the host genome but remain episomal, the expression of the ectopic gene they carry is limited in time and vanishes upon host cell replication. Therefore, the use of this gene expression system in classical gene-therapy approach is restricted only to proof of concept experiments. On the other hand, recombinant adenovirus mediated gene delivery is efﬁcient in inducing developmental redirection since the ectopic gene product serves as a transient trigger to an irreversible process (see below [3]). • Naked plasmid DNA: Over the past several years, signiﬁcant progress has been made in the development of nonviral methodologies that can effectively deliver genes to target tissues in vivo [34]. Delivery of naked plasmid DNA to muscle and liver had a surprising results, showing long term expression of the transgene [35,36].

Potential Sources of Adult Tissues Used for Insulin Gene Therapy Pituitary-Derived Cells For more than 20 years the AtT-20 cell line has been studied as a target for insulin gene therapy. The AtT-20 cell line, derived from the mouse anterior pituitary, synthesizes the adrenocorticotropic hormone (ACTH) precursor, proteolytically processes it to mature ACTH, stores it in secretory granules, and releases mature ACTH upon stimulation with a secretagogue [37,38]. In early studies the cDNA for human proinsulin was introduced into AtT20 cells. The transfected cells, stored immunoreactive insulin, proteolytically

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processed proinsulin into smaller fragments, and secreted the insulin-like material upon stimulation with secretagogues. However, the cells lacked glucose responsiveness [39,40]. Co-infection of both insulin and GLP-1 receptor genes in primary rat pituitary cells using adenovirus-mediated gene transfer resulted in insulin secretion in response to physiological levels of GLP-1 but not of glucose [41]. Co-transfection of insulin with genes encoding the glucose transporter, Glut2, and glucokinase [42,43] resulted in glucose responsive insulin secretion. Transgenic NOD mice in which insulin expression was targeted to the pituitary, showed secretion of biologically active insulin and transplantation of insulin secreting intermediate pituitary lobe tissue reversed diabetes, but insulin secretion was not properly regulated [40]. In conclusion, these studies demonstrate the ability to regulate insulin secretion in pituitary derived cells. However, a major limitation of using these cells for treating diabetes is their continuous secretion of ACTH, which stimulates glucocorticoids synthesis that inhibit insulin action, thus inducing among other things catabolic effects [44]. Intestinal-Derived Cells Neuroendocrine cells from the intestinal epithelium have been suggested as candidates for insulin gene therapy [45–48]. The enteroendocrine cells are found scattered in the intestinal epithelium. These cells synthesize and secrete incretin hormones such as glucose-dependent insulinotropic polypeptide (GIP, from gut K cells) and glucagon-like peptide-1 (GLP-1, from intestinal L cells) after meals, to potentiate insulin production and secretion from the pancreas [49]. Because the secretion patterns of GIP, GLP-1 and insulin are similar and incretins and insulin are compatible in controlling glycemic control, the engineering of enteroendocrine cells for regulated insulin secretion constitutes a feasible approach for insulin gene therapy. K cells transfected with a construct in which human insulin gene expression was controlled by GIP promoter, produced, and secreted insulin in glucose-dependent manner [48]. Glucose sensing ability in these cells is most likely attributed to the glucokinase expression in these cells. Glucokinase is the major glucose-sensing enzyme present also in pancreatic beta cells. Transgenic mice in which the human insulin gene is expressed under the regulation of the GIF promoter [48], produced insulin only in gut K cells, which processed and secreted the biologically active insulin. Gastric insulin production resulted in signiﬁcant amelioration of hyperglycemia in STZ induced diabetic mice for at least 3 months [48]. Additional studies indicated the potential use of L cells as target for insulin gene therapy using rAAV-mediated gene transfer [45]. A genetically engineered human intestinal L-cell line was able to synthesize and secrete

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recombinant insulin. The insulin was secreted via the same granule-like compartments as endogenous GLP-1 and both proteins were secreted with similar dynamic parameters [45]. Since L cells, like pancreatic beta cells, express the endoproteases (PC)1/3 and PC2 [50], the cells processed the proinsulin and secreted the mature insulin in response various stimuli; however it is unclear whether L cells are also glucose responsive. Although promising results from in vitro and in vivo models have demonstrated the potential use of engineered enteroendocrine cells in correcting diabetes [45–48], their use is limited, as the enteroendocrine cells represent only about 1% of the intestinal epithelial population. Therefore, a signiﬁcant challenge still exists in selectively targeting enteroendocrine cells in vivo, or in isolating a pure enteroendocrine population for ex vivo treatment. Gastric-Derived G Cells G cells are endocrine cells located in the stomach. These cells release the peptide hormone gastrin in response to protein-rich meals [51]. G cells synthesize progastrin and process it by PC1/3 and PC2, the same endopeptidases that are implicated in proinsulin processing. Moreover, similar to pancreatic beta cells that store and secrete insulin, these cells store gastrin in secretory vesicles and release it upon proper stimulation [51]. Transgenic mice in which the human insulin gene is expressed under the regulation of the mouse gastrin gene promoter [52], produced insulin only in G cells, which processed and secreted the biologically active insulin in response to meal-associated stimuli. Gastric insulin production resulted in signiﬁcant amelioration of hyperglycemia in STZ-induced diabetic mice [52]. Muscle-Derived Cells Skeletal muscle is a relevant target tissue for gene therapy because of its accessibility and its capacity to produce large amounts of secreted recombinant proteins. Thus, muscle can be considered as a potential target for long-term delivery treatment of chronic diseases [53,54]. In addition, skeletal muscle plays a key role in glucose homeostasis, accounting for up to 70% of glucose disposal after a meal. Numerous groups have studied the use of skeletal muscle in ameliorating diabetes both in vitro and in vivo. Muscle cells, like most non-neuroendocrine cells, lack the speciﬁc endopeptidases (PC2 and PC1/3) required to process proinsulin into active mature insulin. One possibility to achieve proinsulin processing in the absence of the speciﬁc endopeptidases is to engineer a proinsulin molecule that can be correctly cleaved by furin into mature, biologically active insulin. Furin (also known as PACE) is a Golgi-associated propeptide endoprotease that is part of the constitutive secretory pathway of all cells [55].

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The introduction of furin consensus sequences at the B-chain/C-peptide and C-peptide/A-chain junctions has been shown to signiﬁcantly increase the processing of proinsulin to insulin in myoblasts [20,56,57] as well as in other non-neuroendocrine cells such as ﬁbroblasts, epithelial cells, and lymphocytes [58]. In vitro, C2C12 mouse myoblast cells transfected with furincleavable proinsulin cDNAs expressed high levels of insulin mRNA and protein, and about 90% of the proinsulin was processed into mature insulin [20,59]. Engineered myoblast cells ameliorated hyperglycemia upon implantation in diabetic mice [20] and similar results were obtained using primary culture rat myoblasts [22,60]. Skeletal muscle is an accessible tissue which can be efﬁciently targeted using in vivo gene therapy approach. Muscle cells have the ability to take up and express naked plasmid DNA following simple intramuscular injection, avoiding the risks associated with viral vectors. Moreover, it has been reported that despite remaining episomal, the transgene was continuously expressed for up to 19 months following a single intramuscular injection of naked plasmid DNA (reviewed by Ratanamart and Shaw [61]). Intramuscular injection of a plasmid expressing furin-cleavable proinsulin cDNAs into STZ-induced diabetic Balb/c mice resulted in insulin gene expression, production and processing of the hormone. The amelioration of hyperglycemia was associated with increased plasma insulin and C-peptide levels [62–64]. Despite the efﬁcient conversion of furin-cleavable proinsulin into insulin and the promising results using naked plasmid DNA, the efﬁcacy of all the above insulin gene therapy strategies has been severely hindered by the greatly reduced expression and production of furin-cleavable insulin in comparison with the wild-type hormone [62–64]. An additional approach suggested for ameliorating hyperglycemia is to facilitate glucose uptake by peripheral tissues by increasing its transport and its usage [65–68]. Glucose utilization by skeletal muscle is controlled by insulin-stimulated glucose transport through Glut4 and glucose phosphorylation by hexokinase II [65]. Hexokinase II has a low Km for glucose and is inhibited by glucose-6phosphate, which limits glucose uptake. To increase intramuscular glucose usage, the hepatic glucose-phosphorylating enzyme glucokinase (GK) has been expressed in skeletal muscle [66,67]. In contrast to hexokinase II, glucokinase has a high Km for glucose (8 mmol/l) and is not inhibited by glucose6-phosphate [68]. Expression of GK in muscle allows increased glucose uptake by muscle cells in vivo both in transgenic mice [66] and in mice treated by a viral vector to express the transgene [67]. The presence of GK activity in skeletal muscle resulted in increased concentrations of glucose-6-phosphate and glycogen and increased disposal of glucose during an intraperitoneal

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glucose tolerance test. Transgenic mice expressing GK in muscle were also more sensitive to insulin. Recently the skeletal muscle tissue of mice has been engineered to express both furin-cleavable proinsulin and GK transgenes using a single adeno-associated viral vector [69]. Treatment of diabetic mice with this vector restored and maintained normoglycemia under fed and fasted conditions for >4 months with normalization of metabolic parameters assessed by intraperitoneal glucose tolerance testing. Liver Cells Hepatocytes are an attractive target for insulin gene therapy since they have glucose-sensing abilities similar to those in pancreatic beta cells, due to the expression of both Glut2 and glucokinase genes [70]. In addition, the liver is an essential organ that can both store and produce glucose, depending on the relevant physiological state. It has been suggested that even basal hepatic insulin production would have a profound impact on glucose and lipid metabolism in type 1 diabetes. Since hepatocytes, as all the above described nonendocrine cells, lack the speciﬁc endopeptidases function (PC2 and PC1/3), furin cleavable proinsulin analog has been suggested for use in engineering hepatic insulin production [71]. More recently, a single chain analog of insulin has been developed. However, this single-chain insulin exhibited a signiﬁcantly lower biological activity compared to insulin [72] In pancreatic beta cells, glucose controls the level of insulin gene expression, insulin mRNA stability, and the rate of insulin synthesis, processing, and secretion. Such tight regulation by glucose cannot be achieved using liver cells as insulin producers. Attempts to control at least insulin synthesis have focused on utilization of glucose-responsive, liver-speciﬁc promoters. A number of glucose-regulated promoters are available for liver insulin gene therapy [73] and several of them have been studied for glucose-regulated insulin production [24] including glucose-6-phosphatase (G6Pase) whose activity is regulated by glucose. This enzyme catalyzes the hydrolysis of G6Pase functioning in both the gluconeogenic and glycogenolytic pathways. The expression of G6Pase gene is strongly inhibited by insulin [74]. Thus using this promoter may achieve both activation by glucose and inhibition by insulin. Expression of insulin under the control of the G6Pase promoter in the hepatoma cell line HepG2 and in hepatocytes of diabetic rats in vivo resulted, as expected, in glucose-regulated insulin production that was autoregulated by insulin. However, hepatic insulin expression and production was signiﬁcantly lower than pancreatic insulin production [75,76]. The L-type pyruvate kinase promoter [77] and the Spot-14 promoter [78] have also been investigated for their ability to induce glucose-responsive insulin production in the liver. Transcription of the proinsulin gene under the control of these

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promoters was sensitive to glucose levels [79,80]. Injection of recombinant adenovirus or recombinant adeno-associated virus expressing insulin under the control of the L-type pyruvate kinase promoter resulted in nearly normal blood glucose levels in STZ-induced diabetic rats [72,81]. Additional approaches have used promoters that are negatively regulated by insulin to control insulin transgene expression. Including the insulin-like growth factor binding protein-1 basal promoter [81] and the phosphoenolcarboxykinase gene promoter [82]. The use of these promoters to drive insulin gene expression resulted in transient reduction in blood glucose levels after a glucose load, indicating that there is a feedback inhibition on the transcriptional regulation of insulin production [72]. However, the decline in insulin production, exhibited a 3–4 h delay, and more studies are required to develop a system that rapidly responds to changes in blood glucose levels, thus mimicking pancreatic insulin secretion. The glucose transporter Glut2 has been suggested to serve as the actual glucose sensor in the liver [83]. Glut2 expression in the liver is stimulated by glucose and downregulated by insulin [84,85]. These characteristics suggest that the Glut2 promoter may be useful for the regulation of hepatic insulin gene therapy. Therefore, a construct was designed to express the furincleavable human proinsulin gene, under the control of the murine Glut2 promoter. Diabetic mice treated with this construct remained euglycemic for up to 35 days post-treatment. Serum levels of both human insulin and Cpeptide further conﬁrmed the successful transgene delivery and function. These ﬁndings indicate that the Glut2 promoter may be potentially used for driving regulated insulin production for hepatic insulin gene therapy [73]. Despite Glut2 expression in liver, ectopic expression of the Glut2 together with insulin in liver, further increased the glucose-regulated insulin secretion in diabetic rats [86]. Additional systems directed at achieving control of insulin secretion have been based on the use of small molecules. The ability of rapamycin [87] to modulate insulin transgene expression has been analyzed in mouse liver in vivo. A furin-cleavable proinsulin was expressed under the control of a rapamycin-inducible transcription system. Hormone secretion from mouse liver was negligible in the absence of the rapamycin, but was induced in a dose-dependent manner upon its administration, in a process that was reversed following drug withdrawal [87]. In summary, classical gene therapy approaches directed at the production, processing, and glucose-regulated secretion of insulin from extra-pancreatic tissues may carry numerous advantages. This approach promises to replace the diminished insulin production in diabetic patients. Moreover, the extrapancreatic tissues may not become a target for recurrent autoimmune attack which occurs in patients with in Type 1 diabetes. However, the complexity of

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the cellular machinery that allows the proper function of pancreatic beta cells in maintaining continues normoglycemia is extremely difﬁcult to be fully reconstituted in extra-pancreatic tissues, using classical approaches of genetic engineering. For faithfully mimicking pancreatic beta-cell function, the concerted action, and therefore expression, of numerous speciﬁc genes is needed.

Developmental Redirection of Extra-Pancreatic Tissues into Insulin-Producing Cells It has long been thought that animal cells once committed to a speciﬁc lineage, generally, can no longer change their fate and thus become “terminally differentiated” [88,89]. In the last few years, increasing evidence has demonstrated the remarkable ability of some differentiated cells to convert into a completely different phenotype and function [89,90] via a process called developmental redirection or nuclear reprogramming [2,3,89,91]. As opposed to the classical approach of gene therapy, in which a desired function is engineered into a cell by the ectopic expression of the relevant genetic information, developmental redirection involves the guided activation of the relevant endogenous, otherwise silent genetic information in host cells. The process is characterized by wide alterations in the proﬁle of gene expression, allowing adult cells to gain alternate developmental options and therefore functions. The inducing factors were demonstrated to serve as a transient trigger of an irreversible process [2,3]. Several soluble or transcription factors have been identiﬁed to activate the pancreatic lineage in adult extra-pancreatic tissues. The rationale of the approach is that transcription and soluble factors that are known to affect pancreatic organogenesis in the embryo may play similar proactive roles when forced to be expressed in adult nonrelated tissues. PDX-1 was the ﬁrst and most studied pancreatic transcription factor that also possesses the capacity to dictate pancreatic development in differentiated adult tissues. PDX-1 was originally studied because of its central roles in regulating pancreas organogenesis in the embryo and betacell function in adults [92–96]. PDX-1 is involved in regulating the expression of multiple genes in beta cells, including those that encode insulin, glucokinase, islet amyloid polypeptide, PDX-1 itself (in an autoregulatory loop) and, possibly, Glut2 [97]. It plays a key role in pancreatic morphogenesis, as demonstrated by impaired pancreatic development in mice and humans with homozygous mutations in PDX-1 [92–94,98]. In humans, heterozygous mutation of PDX-1 causes maturity-onset diabetes of the young and, probably, type 2 diabetes [99,100]. Heterozygous PDX1-knockout mice have impaired glucose tolerance and an accelerated rate of beta-cell apoptosis [96]. Ectopic

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expression of PDX-1 induced pancreatic lineage and function in BM derived mesenchymal cells, intestinal cells and liver. Bone Marrow as a Pancreatic Progenitor Tissue Bone marrow (BM) cells contain both hematopoietic stem cells and mesenchymal stem cells. BM-derived cells have the ability to differentiate into a number of neuroectodermal, endothelial, mesenchymal, epithelial, and endodermal cell types under special culture condition in vitro or in vivo [101,102]. Additional studies also analyzed the repair roles of BM cells in the restoration of organs function in various diseases including diabetes; these were recently reviewed in refs. [96] and [97]. The capacity of single cell-derived stem cell lines obtained from mouse BM to differentiate into functional insulin-producing cells, has been demonstrated in vitro [103]. The BM derived cells expressed multiple genes related to pancreatic beta-cell development and function including insulin I and II, Glut2, GK, islet amyloid polypeptide, and PDX-1. Insulin and C-peptide were produced and glucose-stimulated insulin release was detected. Moreover, these cells ameliorated hyperglycemia in STZ-induced diabetic mice, without the need for forced expression of pancreatic transcription factors [103]. Multiple studies analyzed whether BM transplantation may results in betacell restoration and normoglycemia. The outcomes were differing, in some studies BM-derived cells could differentiate into beta cells, whereas others show, at best, mixed results and at worst none (101–106). Whereas Ianus et al. [104] presented evidence that BM-derived cells can differentiate into beta cells and ameliorate diabetes upon implantation into irradiated mice, other groups failed to reproduce these results and suggested that the donor BM-derived cells may have promoted the endogenous beta-cell expansion [105,106]. Transplantation of BM into STZ-induced diabetic mice may have initiated proliferation of the residual pancreatic cells resulting in improved control of the blood glucose levels. Although some insulin-positive BM-derived cells were present in sections of STZ-damaged pancreatic tissue, their low frequency, and the lack of the transcription factor PDX-1 expression, suggest that the BM-derived cells did not, by themselves, functionally rescue the recipients [105]. It was suggested that BM-derived cells are recruited to the pancreas in response to islet injury, and by a yet unknown mechanism, these cells induce endogenous pancreatic tissue repair. The BM-derived cells may facilitate the recovery of nonterminally injured beta cells or may improve the survival and/or function of islets by inducing neo-vascularization of the pancreas [106]. The capacity of pancreatic transcription factors to route BM derived cells differentiation towards beta cells have been recently analyzed. It was shown

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that ectopic expression of PDX-1 activated the pancreatic lineage and function in BM derived mesenchymal cells isolated from human donors [107]. RNA and protein analyses documented the activation of expression of all four islet hormones. A signiﬁcant insulin content, as well as glucose-stimulated insulin release, were demonstrated in vitro. Cell transplantation into streptozotocin-diabetic immunodeﬁcient mice resulted in further beta-cell differentiation and the amelioration of hyperglycemia [107]. Differentiation of Immature Intestinal Cells into Insulin-Producing Cells The small intestinal epithelium includes a large population of undifferentiated cells. These cells, which are present in the crypt region, are known to serve as stem cells and differentiate into intestinal epithelium cells and endocrine cells. Since these adult organ-speciﬁc stem cells have retained developmental plasticity, their ability to differentiate into insulin-producing cells was analyzed [108,109]. The IEC-6 immature intestinal crypt cell line was used for these studies. IEC-6 cells are derived from the small intestine of a normal rat, and have an undifferentiated morphology and limited expression of intestinal speciﬁc genes. Ectopic expression of the pancreatic transcription factor PDX1 enabled the IEC-6 cells to express multiple pancreatic speciﬁc genes such as amylin, glucokinase, and Nkx6.1, originally not expressed in these cells. However, insulin gene expression was not detected in PDX-1–transfected IEC-6 cells [108]. Co-treatment of PDX-1 with betacellulin [108] or the combined expression of PDX-1 and Isl-1 [109] was required for the production of insulin in IEC-6 cells. Although insulin accumulated in secretory granule-like structures, secretion was not sensitive to glucose. Implantation of the PDX-1 treated cells into diabetic rats reduced plasma glucose levels [108,109]. This study demonstrates the instrumental role of pancreatic transcription factors in dictating differentiation toward the pancreatic lineage. However, IEC-6 cells maintain high levels of plasticity, possibly due to the fact that these cells are not terminally differentiated. Liver to Pancreas Developmental Redirection The liver is the largest organ in our body with a high level of functional redundancy [110]. Unlike beta cells, the liver regenerates efﬁciently, mainly by the proliferation of mature hepatocytes [111]. Human liver cells can be propagated in vitro for months, and the numbers of cells can be expanded substantially ex vivo. Trans-conversion between liver and pancreas is conceivable since liver and pancreas are developmentally related, as both are derived from appendages of the upper primitive foregut endoderm. It has been suggested that the late separation of liver and pancreas during organogenesis in primitive ventral endoderm might have left both tissues with pluripotent cells that are capable of giving rise to both hepatic and pancreatic

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lineages [112]. Both tissues have many characteristics in common, including responsiveness to glucose and a large group of speciﬁc transcription factors mutually expressed in both tissues [113]. Trans-conversion between pancreatic acinar cells and hepatocytes in both rodents and humans has been reported under experimental, pathologic and malignant conditions [114]. Comparing the development of liver and pancreas in other species further emphasizes the close relationship between these two organs. In lower organisms, such as worms and ﬁsh, there is no spatial separation between the two organs [115,116] and the “hepato-pancreas” functions as both liver and pancreas [112]. Liver to Pancreas Developmental Redirection In Vitro Using embryonic or fetal liver cells. It is reasonable to assume that immature liver cells may be more susceptible to acquire pancreatic characteristics due to their pluripotency. The bile duct-derived progenitors called oval cells have been characterized in rodents, but their human equivalents have not yet been found [111]. These cells were documented previously to possess several developmental options, including that of hepatocytes, biliary epithelium, intestinal epithelium and pancreatic acinar epithelium [117]. Yang et al. [118] demonstrated that exposure to high levels of glucose and nicotinamide in vitro causes oval cells to acquire an additional developmental option: that of insulin-producing cells. These cells express several genes that encode pancreatic transcription factors, including PDX-1 [118]. The induction of a functional pancreatic lineage has also been documented in telomerase expressing fetal human liver cells, treated with a lentivirus to stably express PDX-1 [119]. Upon treatment with soluble factors such as activin-A and serum withdrawal, these cells exhibited improved pancreatic beta-cell-like function, such that insulin production in these cells was about 60% of that of normal human pancreatic beta cells [120]. Processed insulin was secreted in a glucose regulated manner and the cells ameliorated hyperglycemia in diabetic SCID-NOD mice for long periods [119, 120]. Using adult liver cells. The capacity of inducing functional redirection of mature liver cells carries a substantial therapeutic signiﬁcance, since it may allow autologous cell replacement therapy for diabetics. Indeed, a recent study demonstrated the potential use of primary cultures of adult human liver cells as a source of pancreatic progenitors [121]. Cells isolated from adult human liver were propagated in vitro for many passages, and upon expression of PDX-1 and treatment with EGF and nicotinamide, up to 50% of PDXexpressed some beta-cell speciﬁc characteristics. Moreover, the cells produced insulin, correctly processed and stored it in induced secretory granules and secreted the hormone upon glucose challenge [121]. The most important “pancreatic-like” function these cells demonstrated was their capacity to

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ameliorate hyperglycemia upon implantation in diabetic immunodeﬁcient mice in vivo. Human C-peptide secretion and the parallel amelioration of diabetes persisted for the whole duration of the experiment (60 days). Consistent with the functional properties of PDX-1 treated adult human liver cells was the expression of the pancreatic transcription factors, including that of the endogenous human PDX-1 [121]. Li and collaborators [90] documented that in PDX-1-treated HepG2 cells as well, the transgene was only a temporary trigger that was no longer required once the pancreatic differentiation program was activated. A Summary Of The Main Approaches Using Adult Liver Cells In Vitro Is Presented In Table 2a.

Table 2. Activation of pancreatic lineage in liver by developmental redirection process Study Cells/Species Treatment Result a. In vitro studies Sapir et al. [121]

Primary culture of adult human liver cells

PDX-1+Nicotinamide +EGF

Pancreatic speciﬁc gene expression Insulin production and secretion Amelioration of hyperglycemia upon transplantation in diabetic mice

Zalzman et al. [119,120]

Telomerase expressing fetal human liver cells

PDX-1+Activin A

Pancreatic speciﬁc gene expression Insulin production and secretion Amelioration of hyperglycemia upon transplantation in diabetic mice

Coa et al. [142]

Rat WB-1 cells

PDX-1–VP16

Partial pancreatic speciﬁc gene expression Maturation upon transplantation in diabetic mice

Nakajima-Nagata et al. [143]

Rat small hepatocytes

PDX-1

Pancreatic speciﬁc gene expression Insulin production and secretion

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Table 2. Continued Study

Cells/Species

Treatment

Result

Human hepatoma cells, HepG2

PDX-1–VP16

Pancreatic speciﬁc gene expression Insulin production and secretion

b. In vivo studies Ferber et al. [124,125,141]

Mice

PDX-1

Pancreatic speciﬁc gene expression Insulin production and secretion Amelioration of hyperglycemia

Horb et al. [123]

Xenopus tadpoles

PDX-1–VP16

Pancreatic speciﬁc gene expression

Imai et al. [136]

Mice

PDX-1–VP16

Insulin production and secretion Amelioration of hyperglycemia

Kaneto et al. [134,135]

Mice

PDX-1–VP16/MafA/ NeuroD1/Ngn3

Insulin production and secretion Amelioration of hyperglycemia

Koizumi et al. [140]

Mice (40% hepatectomy)

PDX-1

Pancreatic speciﬁc gene expression Insulin production and secretion Amelioration of hyperglycemia

Kojima et al. [122]

Mice

NeuroD1/betacellulin

Pancreatic speciﬁc gene expression Insulin production and secretion Amelioration of hyperglycemia

Li et al. [144]

Developmental Redirection of Liver to Pancreas In Vivo Studies, performed both in mice and frogs (Xenopus), suggest that PDX-1 and other pancreatic transcription factors activate the pancreatic lineage in liver also in vivo, via a process called transdifferentiation [122–124]. Transdifferentiation describes an irreversible switch of one type of differentiated cell into another differentiated cell [89]. Transient, ectopic expression of PDX-1 in liver, delivered by systemic administration of ﬁrst-generation E1-deleted recombinant adenovirus (FGAD), induced a wide repertoire of pancreatic gene expression. Sur-

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prisingly the ectopic PDX-1 expression led to long-lasting production and secretion of processed, biologically active insulin, despite the short term expression of PDX-1 transgene in liver [124,125]. PDX-1 induced its own expression in liver (auto-induction) along with the expression of numerous pancreatic transcription factors, which, in turn, explains the irreversible nature of the “liver-to-pancreas” developmental redirection process [124]. Hepatic insulin production triggered by PDX-1 administration using FGAD was functional. It not only restored euglycemia in STZ-induced diabetic mice [125], but also prevented STZ-induced hyperglycemia, even eight months after the initial FGAD treatment [124]. These data indicate the irreversibility of the process and that the insulin-producing cells in liver resist beta-cellspeciﬁc toxins. Moreover, since the liver plays an important role in neutralizing toxins and, in contrast to pancreatic beta cells, has high levels of catalase and superoxide dismutase activities [110], the surrogate beta cells derived from liver may resist cellular assaults to which beta cells are sensitive. When ectopically expressed in the mature, fully differentiated liver, PDX-1 functions as a pancreatic differentiation factor, having the capacity to induce the expression of multiple pancreatic genes, including several that are not considered its immediate targets, such as glucagon, p48, and elastase [124]. PDX-1-induced liver to pancreas transdifferentiation by the FGAD delivery system occurred in predisposed cells, comprising less than 1% of cells in the liver located mainly close to the central veins, despite the initial random expression of the transgene in liver. Such a predisposition could be caused by the heterogeneity of cells in liver [126]. Different cells could respond distinctly to the pancreatic transgene in the sense of ultrastructural modiﬁcations in chromatin compaction, presence of silencing effects, or lack of complementing transcription factors that may work in concert with the ectopic gene [127–130]. Partially Overcoming the Hepatic Predisposition To overcome some of the barriers cells may have to the PDX-1 induced transdifferentiation process, and for increasing the number of “responding” cells, Horb et al. [123] expressed the PDX-1 as a fusion protein with the transcriptional activation domain of VP-16 (Xlhbox8–VP16), a regulatory protein from herpes simplex virus. PDX-1–VP-16 expression in transgenic tadpoles induced the conversion of most of the liver into exocrine and endocrine pancreas. The massive transformation of the tadpole’s liver into pancreas, suggests that activation of PDX-1 by the VP-16, might have eliminated the requirement for additional co-factors thought to work in concert with PDX-1 in activating pancreatic gene expression [131]. Alternately, VP-16 could also affect chromatin condensation and remodeling of PDX-1 target genes of the host genome [132,133]. Activation of PDX-1 may have overcome some of the

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barriers of hepatic cell predisposition, thus, strengthening the notion that this predisposition is caused in part by complementing, pre-existing transcription factors, and also possibly by distinct chromatin structure. However, PDX-1–VP-16 had only moderate effects on the activation of pancreatic lineage when ectopically expressed in adult mice liver, compared to its effects in Xenopus. While the VP-16-activated PDX-1 was more efﬁcient than the wtPDX-1 in ameliorating hyperglycemia, in mice, only a fraction of adult liver cells acquired the pancreatic characteristics. These differences could be explained by distinct responses of amphibians and mammals to the ectopic gene or could be attributed to the distinct embryonic differentiation stage of the treated hepatic tissues; fetal tissue (Xenopus) could be more permissive to undergo a developmental redirection process than mature mouse liver [134–136]. Several lines of evidence taken together with the massive liver transformation suggest that PDX-1 induces in liver a transdifferentiation event: ﬁrst, the Xlhbox8–VP16 transgene driven by the transthyretin promoter (a gene usually expressed in mature hepatocytes) was not expressed until a day after the liver bud had begun expressing hepatic markers [123]; second, the PDX-1 mediated process involves the down-regulation of hepatic markers expression [123]. Similar effects of PDX-1 on the hepatic repertoire of gene expression has been demonstrated also in mice [137] Additional Pancreatic Transcription Factors Induce Pancreatic Lineage in Liver Several pancreatic transcription factors have been shown to promote the PDX-1 effect on the developmental redirection process, preferentially betacell maturation, as manifested by improved amelioration of hyperglycemia. While NeuroD1, MafA or Ngn3 were incapable of individually inducing betacell function in liver, they signiﬁcantly promoted the PDX-1 effect on the process [134,135]. In one study, NeuroD1 expression alone moderately activated the pancreatic repertoire and function in mice, but its effect was substantially augmented when combined with betacellulin administration [122]. During pancreas development, this basic helix-loop-helix transcription factor is essential for islets morphogenesis and proper function [138]. Interestingly, ectopic NeuroD1 expression in liver induced both downstream and upstream transcription factors that are part of the pancreatic transcriptional network, including PDX-1. However, while in PDX-1-treated liver, glucagon and insulin were produced in distinct cells [124], all the pancreatic hormones were coproduced within the same cell in mice liver treated with NeuroD1 [122]. The appearance of distinct types of pancreatic hormone-producing cells may suggest that the developmental shift upon expression of PDX-1 occurred in distinct populations of predisposed liver cells [124]. It is not clear what

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dictates the distinct characteristics of the cells induced: the distinct transcription factors; the distinctly affected host cells in liver; or possibly both. Soluble Factors Promote Liver-to-Pancreas Transconversion Process A role for soluble factors in promoting transcription factors-induced liver to pancreas transdifferentiation has been suggested. Betacellulin (beta-cellstimulating hormone), a member of the epidermal growth factor family of growth factors [139], has been implicated in promoting NeuroD1-induced liver to pancreas transdifferentiation in mice [122]. Combined ectopic expression of NeuroD1 and betacellulin induced higher levels of insulin production and ameliorated hyperglycemia in STZ diabetic mice in a glucoseregulated manner. Hepatic regeneration using 70% hepatectomy was also demonstrated to improve the effect of PDX-1 in accelerating the transdifferentiation process along the pancreatic lineage [140]. The improved therapeutic outcome of the process could be due to the increased number of proliferating and possibly pluripotent cells in this organ. The effect of partial hepatectomy on PDX1-induced transdifferentiation could be possibly mimicked by co-treating the mice with growth factors such as hepatic growth factor or interleukin-6, the levels of which increase during liver regeneration. A Summary Of The Main Approaches Using Adult Liver Cells In Vivo Is Presented In Table 2b.

Next Steps in the Developmental Redirection Paradigm Additional efﬁcacy studies should indicate whether developmentally shifted cell function persists over long periods, as type 1 diabetes in humans is a multi-decade disease. Safety studies need to be performed to determine whether these cells may contribute to inappropriate beta-cell hyperplasia. Will these cells lead to dangerous hypoglycemia? Can these cells be delivered to the liver, as in islets cell implantations in humans? Will insulin-producing cells revert into hepatocytes if introduced back into the liver? Since the beta cells that are created with these methods produce relatively small quantities of insulin, can they be induced to produce higher levels of the hormone to mimic normal human islets or even become super-islets? Can insulin be secreted within 2 or 3 min of exposure to glucose, as occurs in normal beta cells? Will the developmentally shifted mature extra-pancreatic cells become a target for autoimmune attack that characterizes type 1 diabetes? Importantly, a recent study demonstrates that ectopic PDX-1 expression in liver in vivo ameliorated diabetes also in the autoimmune model of diabetes; the Non-Obese-Diabetic mouse (NOD). Interestingly, along with the amelioration of hyperglycemia, the PDX-1-treated mice also exhibited an immune

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modulation process which was associated with a decrease in the Th1 response and an increase in the Th2 response to diabetogenic antigens [141].

Conclusion Reconstruction of beta-cell function in extra-pancreatic tissues may constitute a substantial stride toward the development of cell-replacement therapy for diabetic patients. The usage of adult extra-pancreatic cells for generating functional insulin-producing tissue may pave the way for autologous implantations, thus allowing the diabetic patient to be the donor of his or her own insulin-producing tissue. This approach may circumvent the shortage in tissue availability, the need for anti-rejection treatment, and the ethical issues associated with the use of fetal or embryonic stem cells for this purpose. However, it is obvious that the generation of a cell that will faithfully mimic the pancreatic beta cell requires the concerted action of many factors which are uniquely expressed only in pancreatic beta cells. While classical gene therapy may allow the expression of a few genes at once and thereby only partially restoring some of the beta-cell characteristics, a process of nuclear reprogramming or developmental redirection may allow the generation of a comprehensive beta-cell phenotype and function. The studies reviewed in this chapter serve as a proof of concept for the basic capacity of directing the differentiation of cells in extra-pancreatic tissues toward the pancreatic lineage and function. A further “ﬁne-tuning” of the combination of transcription and soluble factors is expected to increase the therapeutic merit of this novel regenerative medicine approach, called nuclear reprogramming. Adult tissues, such as BM-derived mesenchymal cells and liver cells, may retain substantial plasticity and can be induced to assume new fates and function, upon appropriate molecular manipulation. Unraveling the mechanism(s) of such a process may allow the use of additional adult tissues in generating beta cells as well as other cell types, so that we can exploit our own adult tissues to generate organs in need. Acknowledgments. We thank the members of our laboratory who contributed to the studies discussed here: Keren Shternhall, Tamar Sapir, Shiraz Geffen-Halevy, Odelia Nakar, and Dana Berneman-Zeitoni. We also thank Itzick Rachmut, Vered Elbaz-Aviv, Kﬁr Molakandov, and Michal Mauda for fruitful discussions and for critically reviewing the manuscript.

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IV. Incretins and Beta-Cell Function

16. Molecular Biology of Gluco-Incretin Function Sonia Klinger and Bernard Thorens

Summary. The gluco-incretin hormones glucose-dependent insulinotropic polypeptide (GIP) and glucagon like peptide-1 (GLP-1) are secreted by intestinal endocrine cells and have been studied for many years because of their important effect to potentiate glucose-stimulated insulin secretion. In contrast to GIP, GLP-1 retains its insulinotropic effect in type 2 diabetic patients and a long-acting agonist of this peptide is now used for the treatment of this disease. Both peptides, however, have also long-term beneﬁcial effect on the preservation or augmentation of the pancreatic beta-cell functional mass by stimulating beta-cell differentiation from precursors, proliferation of mature beta cells, and their protection against apoptosis. Although several studies have investigated the underlying molecular mechanisms, much remains to be learned about the mode of action of these hormones on beta cells. Here we review the current knowledge on gluco-incretin biology with a speciﬁc perspective on their beta-cell action. We also discuss the role that GLP-1 has on beta cells through indirect mechanisms, in particular through the regulation of the hepatoportal vein glucose sensors. This serves as a reminder that betacell function needs to be also studied in an integrated physiological approach where the complexity of their functional regulation can be appreciated.

Introduction The absorption of food triggers multiple endocrine and nervous responses that are required to coordinate nutrient storage in different organs and to regulate feeding termination. The gut is the site of production and release of many such hormones, a subclass of them have the additional role of potentiating the effect of glucose on insulin secretion. These hormones, referred to Department of Physiology and Center for Integrative Genomics, University of Lausanne, Genopode Building, Lausanne, Switzerland

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as gluco-incretins, are glucagon-like peptide-1 (GLP-1) and glucosedependent insulinotropic polypeptide (GIP). Because GLP-1 has a conserved capacity to potentiate insulin secretion in a glucose-dependent manner even in type 2 diabetic patients—a property not observed for GIP—it has been extensively studied for its potential therapeutic utilization for the treatment of type 2 diabetes. GLP-1, long-acting analogues of this hormone, or inhibitors of the enzyme that degrades it are already in the clinic or being evaluated in clinical trials. However, much remains to be learned of the physiology of GLP-1 and GIP. In particular, besides the role of GLP-1 in stimulating insulin secretion, this peptide and also GIP have long-term actions in stimulating beta-cell differentiation from precursor cells, in inducing mature beta-cell proliferation and their protection against stress-induced apoptosis. These effects combine to regulate beta-cell mass and function, and may be critical to improve insulin secretion in type 2 diabetes. In addition, GIP and GLP-1 have extrapancreatic effects which regulate many physiological functions related to glucose and energy homeostasis. Interestingly, GLP-1 can also be produced by neurons of the brain stem which projects towards hypothalamic nuclei and probably also higher brain structures such as the hippocampus, where GLP-1 receptors are located and may modulate memory acquisition and retention. Thus, although GIP, and GLP-1 in particular, are becoming important therapeutic tools, their complex physiological functions are still incompletely understood. Uncovering their mechanism of action may lead to the discovery of important new regulatory mechanisms.

GLP-1 and GIP Structure and Biosynthesis The preproglucagon molecule is synthesized in pancreatic alpha-cells, in gut endocrine L cells located in the jejunum and colon, and in neurons present in the brainstem, in particular in the nucleus of the solitary tract [1,2]. Preproglucagon is a 180-amino-acid long prohormone and its sequence contains, in addition to glucagon, two sequences of related structure: GLP-1 and GLP-2 (Fig. 1). Alpha-cell-speciﬁc proteolytic processing of preproglucagon [3,4] leads to the production of glucagon as the major biologically active peptide (Fig. 1). In intestinal L cells, preproglucagon is proteolytically cleaved to generate glicentin and oxyntomodulin, two peptides with unclear physiological functions at the concentrations they are found in the plasma, and GLP-1 and GLP-2, but not glucagon (Fig. 1). The brainstem neurons produce both GLP-1 and GLP-2. Prohormone convertase PC1/3 is essential for proglucagon processing in L cells [5], and PC2 is critical for the generation of glucagon in pancreatic alpha-cells [6]. Whereas GLP-1 produced by L cells is released in the blood as an endocrine hormone, GLP-1 produced by brainstem neurons

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glicentin oxyntomodulin GLP-1 GLP-2 IP-2

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Fig. 1A,B. Glucagon-like peptide-1 (GLP-1) and glucose-dependent insulinotropic polypeptide (GIP) precursors. A Structure of the mammalian proglucagon precursor and the proglucagon-derived peptides generated in different tissues. B Structure of the mammalian GIP precursor. The upper numbers correspond to the rat sequence, the lower numbers correspond to the human sequence

most probably acts as neurotransmitter in a central nervous system-restricted GLP-1/GLP-1 receptor signaling pathway. GIP is a 42-amino-acid long hormone produced from preproGIP in intestinal K cells (Fig. 1) [7]. The secretion of both GLP-1 and GIP can be stimulated by absorbed glucose and fat. In the postprandial state, however, increase in circulating GLP-1 and GIP occurs with the same kinetics even though K cells are located in the duodenum and L cells are located in the distal intestine and colon. Therefore, GLP-1 release may be stimulated by neurotransmitters from the enteric nervous system, or by hormones released from cells of the proximal intestine, such as GIP itself [8].

GLP-1 and GIP Receptors The structure of the GLP-1 receptor was ﬁrst elucidated from the structure of its cDNA isolated from a rat islet cDNA expression library [9]. The human islet receptor has also been cloned [10], and identical receptor cDNAs were also isolated from rat lung and hypothalamus [11]. The GLP-1 receptor belongs to the same family of G-protein-coupled receptors as those for glucagon, GIP, vasoactive intestinal polypeptide (VIP), pituitary adenylyl cyclase

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activating polypeptide (PACAP), and several others [12]. The amino-terminal extracellular domains of these receptors are relatively long with ∼120 amino acids, and they do not share any signiﬁcant sequence homology except for the presence of six cysteines, which have been perfectly conserved at identical locations in all the receptors of this family. Binding of GLP-1 to its receptor is with high afﬁnity (Kd ∼0.2 nM) and speciﬁcity. Indeed, peptides of related structures such as GIP or VIP do not bind the receptor [9]. Interestingly, exendin-4 and its truncated form exendin-(9–39), isolated from the venom of the Gila monster Heloderma suspectum, a venomous lizard found in the United States and Mexico, bind to the human and rat receptors with high afﬁnity [10,13], with exendin-4 being a full agonist and exendin-(9–39) an antagonist. The GIP receptor cDNA was originally cloned from a rat brain cortex cDNA library [14]. It encodes a seven-transmembrane domain protein consisting of 455 amino acids. The human [15,16] and rat [17] receptors were also cloned and encode proteins of, respectively, 466 and 462 amino acids. The GIP receptor binds its ligand with a Kd of ∼0.2 nM [15]. GIP(6–30)–NH2 binds to the receptor with a similar afﬁnity, but acts as an antagonist, inhibiting GIPinduced cAMP formation [18]. GIP-(7–30)–NH2 is a potent antagonist which can markedly reduce glucose-stimulated insulin secretion in fasted rats, or blunt the postprandial increase in plasma insulin levels [19]. Exendin-(9–39) is also an inhibitor of GIP binding and of cAMP formation by GIP-activated receptors [15].

Intracellular Signaling by the GLP-1 Receptor The multiple actions of gluco-incretin hormones on differentiation of betacells from precursor cells, replication of mature beta-cells, or their protection against apoptosis probably require activation, or inhibition, of diverse transcriptional programs (Fig. 2). The signaling pathways activated by GLP-1 in beta-cells depend principally on the initial production of cAMP [20]. Protein kinase A (PKA)-mediated phosphorylation of the transcription factor cAMP responsive element binding protein (CREB) is the classical mode of transcriptional regulation by GLP-1. This pathway is also known for glucose-mediated beta-cell survival [21]. GLP1 also activates the extracellular signal-regulated kinase 1/2 (ERK1/2) pathway [22,23], probably through a combination of increased cAMP and [Ca2+]i activating the protein kinase Raf [24,25]. The activation of the phosphatidylinositol-3-kinase (PI3K) pathway has also been documented and could be either through a direct activation by the beta/gamma subunits of Gs [26], or through an indirect pathway involving c-src activation of betacellulin and the

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Fig. 2. Gluco-incretin hormone signaling pathways controlling gene transcription. Binding of the gluco-incretin hormones to their receptors stimulates, via the Galphas subunit, adenylyl cyclase (AC), resulting in the production of cyclic AMP (cAMP). cAMP activates protein kinase A (PKA) which phosphorylates the transcription factor cAMP responsive element binding protein (CREB). Gene transcription can also involve Ca2+ release from the endoplasmic reticulum (ER) via cAMP-Epac2-mediated sensitization of the ryanodine receptor. This event requires extracellular Ca2+ inﬂux through voltagedependent Ca2+ channels (VDCC) activated by the glucose signaling pathway. Ca2+ released from the ER activates extracellular-signal regulated kinase 1/2 (ERK1/2) signaling cascade by an undeﬁned mechanism. The third pathway initiated by GIP and GLP-1 activates phosphatidylinositol-3-kinase (PI3K), either directly through the beta/gamma subunits of G-protein, or indirectly through the c-src-mediated activation of the epidermal growth factor (EGF) receptor. The PI3K/Akt pathway can also be activated by activated insulin receptor substate-2 (IRS-2), whose expression is under the transcriptional regulation by the cAMP/CREB pathway and which is downstream of the insulin and/or insulin growth factor-1 (IGF-1) receptors. One of the mechanisms of action of Akt is to inactivate Foxo1 by phosphorylation, thereby inducing its nuclear exclusion. Active Foxo1 inhibits the transcription of PDX-1 and its translocation to the nucleus

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epidermal growth factor (EGF) receptor [27] and possibly implicating PKCzeta [28].

GIP, GLP-1, and the Potentiation of Glucose-Stimulated Insulin Secretion One of the most studied effects of GIP and GLP-1 is their action to potentiate glucose-stimulated insulin secretion [29–32], an effect that requires glucose to be present at least equal to normoglycemia and which increases the magnitude of the insulin secretion bursts without modifying their frequency [33]. Glucose induces insulin secretion by activating a metabolism-dependent signaling pathway that is initiated by glucose entry into the beta cells through the glucose transporter GLUT2, and its phosphorylation by glucokinase. This enzyme catalyzes the rate-controlling step in glucose metabolism and mitochondria-derived factors, in particular an increase in the ATP/ADP ratio, which leads to closure of KATP-dependent K+ channels, membrane depolarization and inﬂux of Ca2+ through voltage-dependent channels. The inﬂux of Ca2+ eventually triggers insulin exocytosis. The stimulatory effect of the gluco-incretin hormones on the glucose signaling pathway requires binding of these hormones to their speciﬁc beta-cell surface receptors. These receptors are linked to activation of the cAMP pathway [9,14] and the mechanisms responsible for ampliﬁcation of the glucose signal inducing insulin secretion involve both a PKA-dependent and a PKA-independent pathway. Activated PKA can phosphorylate several proteins involved in the sensing of glucose and modify their activity. This is the case for GLUT2, the KATP channel, and the L-type Ca2+ channel. Phosphorylation of GLUT2 reduces its Vmax for glucose transport [34], and phosphorylation of the KATP channel reduces its sensitivity to changes in glucose concentrations [35]. Thus, both of these actions would tend to reduce rather than augment glucose-stimulated insulin secretion and may therefore control the termination of the glucose signal. Phosphorylation of the Ca2+ channel, however, increases its sensitivity to plasma membrane depolarization. Another apparently important mechanism of action of cAMP/PKA, which is synergized by activated PI3K, is to reduce the sensitivity of the voltagedependent K+ channel Kv2.1 to changes in plasma membrane depolarization. This prolongs the duration of action potentials of the beta-cells and thus contributes to increase glucose-stimulated insulin secretion [36,37]. One still not completely elucidated site of action of PKA is the exocytotic step, where electrophysiological studies have indicated an important regulatory role for cAMP [38] (Fig. 3).

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Fig. 3. Gluco-incretin hormone signaling pathways controlling glucose-stimulated insulin secretion. Glucose induces insulin secretion by activating a metabolism-dependent signaling pathway that is initiated by glucose entry into the beta-cells through GLUT2. Glucose metabolism leads to an increase in the ATP/ADP ratio, which induces the closure of ATPdependent K+ channels. This leads to membrane depolarization (∆Ψ) and inﬂux of Ca2+ through voltage-dependent calcium channels (VDCC), which triggers insulin granule exocytosis. The effects of the gluco-incretin hormones in this process are diverse. First, they activate protein kinase A (PKA), which phosphorylates GLUT2, the KATP channel, the Ca2+ channel, the voltage dependent K+ channels (Kv2.1), and regulatory proteins involved in insulin exocytosis. Second, they activate, in a PKA-independent manner, Epac2, a cAMP sensor protein. Epac2 forms a complex with Rim2 and interacts with Rab3A to stimulate insulin exocytosis. Epac2 also promotes Ca2+ release from the endoplasmic reticulum by sensitizing the ryanodine receptor

The PKA-independent pathway involves activation of the cAMP sensor molecule cAMP-regulated guanine nucleotide exchange factor II (cAMPGEFII) or Epac2 which forms a complex with Rim2 and interacts with Rab3A to stimulate insulin granule exocytosis [39]. Interestingly, cAMP binds to cAMP-GEFII with a Kd in the micromolar range whereas its afﬁnity for the regulatory subunit of PKA is ∼50 nM [40]. Thus, relatively low intracellular cAMP levels are sufﬁcient to induce the phosphorylation of speciﬁc PKA targets whereas higher cAMP levels are needed to recruit the cAMP-GEFII pathway. Epac2 has also been involved in the control by cAMP of the ryanodine receptor which acts as a Ca2+-activated Ca2+ channel leading to release

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in the cytoplasm of Ca2+ stored in the endoplasmic reticulum (ER) [41] (Fig. 3).

Gluco-Incretin Hormones and Beta-Cell Differentiation from Precursors and Proliferation of Mature Beta-Cells In recent years, both GIP and GLP-1 have been shown to have trophic effects on pancreatic beta-cells. These are evidenced, in part, by their effect on the differentiation of precursors into insulin-secreting cells. For instance, the pancreatic tumor cells AR42J, which have mixed exocrine and endocrine characteristics, progressively acquire the capacity to synthesize insulin, and to express GLUT2 and glucokinase when cultured in the presence of GLP-1 [42]. Similarly, exendin-4 treatment of Capan1 cells derived from pancreatic duct cells activates the expression of GLUT2, glucokinase but also the transcription factors pancreatic and duodenal homeobox gene-1 (PDX-1), Beta2/ NeuroD and hepatocyte nuclear factor 3beta (HNF3beta/Foxa2), which are necessary for the beta-cell phenotype [43]. Nestin-positive, islet-derived precursor cells expressing the GLP-1 receptor, can be stimulated by GLP-1 to differentiate into insulin-secreting, PDX-1-expressing cells [44]. PDX-1 appears essential for GLP-1 to induce beta-cell differentiation, since ductal PANC-1 cells fail to differentiate into insulin-producing cells in response to GLP-1 in the absence of PDX-1 expression [45]. The precise mechanisms involved in GLP-1-dependent beta-cell differentiation are only poorly deﬁned, but may require activation of PKC and mitogen-activated protein kinase (MAPK) [42], as well as increased insulin receptor substrate-2 (IRS-2) expression and translocation of forkhead box protein O1 (Foxo1) from the nucleus to the cytoplasm [46]. In vivo, GLP-1 or the long-acting agonist exendin-4, has been shown to both increase beta-cell mass and improve glucose-stimulated insulin secretion in various experimental conditions. For instance, daily injection of exendin-4 to pancreatectomized rats for 10 days induces a signiﬁcant increase in beta-cell mass [47], an effect associated with an increase in PDX-1 expression [48]. Also, administration of exendin-4 to db/db mice signiﬁcantly improves glycemia, with a concomitant increase in insulinemia and reduction in glycated hemoglobin HbA1c [48]. A very interesting observation was that rats injected with streptozotocin at the time of birth, a treatment which induces a form of type 2 diabetes at 2 months of age, were protected from development of diabetes when injected daily with GLP-1 or exendin-4 between day 2 and day 6 after birth [49]. This was associated with a relative increase in beta-cell mass and pancreatic insulin content as compared to streptozotocin-treated rats. However, the secretory response of the glucose-

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perfused pancreas was not corrected, but hepatic glucose production was signiﬁcantly decreased. Thus, in addition to an insulinotropic action, the early treatment with GLP-1 or exendin-4 may also induce changes in the control of liver glucose metabolism, which may be due to changes in its autonomic innervation. A similar observation was reported by the same authors concerning the prevention of diabetes in the genetically diabetic GK rat [50]. In a rat model of intrauterine growth retardation, which leads to development of a diabetic phenotype at adult age, administration of exendin-4 during the prediabetic neonatal period prevents the development of diabetes. This is due to the complete prevention of beta-cell number reduction that is normally observed in the retarded rats, probably through the preserved expression of PDX-1 and normalization of islet beta-cell proliferation rates [51]. GLP-1 induction of beta-cell proliferation has been initially evaluated by Buteau and colleagues using the INS-1 and INS(832/13) clonal cell lines and conﬁrmed in rat islet cells [27,28,52]. These authors showed that GLP-1 induced incorporation of [3H]thymidine in these cells through a PI3K-dependent mechanism. They identiﬁed PKCzeta, Akt/PKB, and MAPK as potential downstream effectors of PI3K. Increased expression of Akt/PKB and MAPK/ ERK has also been observed in 8-week-old db/db mice treated with exendin-4 [53]. They further proposed that this proliferation mechanism may involve c-src-mediated increase in betacellulin cleavage and activation of the EGF receptor. Whether this is the sole mechanism by which GLP-1 can activate beta-cell replication is not known. In addition, it should be noted that the proliferative action of GLP-1 is rather modest, indicating that only a subfraction of beta-cells may be susceptible to its action and able to go through a complete cell cycle.

GLP-1 and Protection Against Apoptosis Apoptosis is a critical mechanism leading to reduction of beta-cell mass in both type 1 and type 2 diabetes. Whereas type 1 diabetes is induced by autoimmune and inﬂammatory reactions [54,55], reduction in beta-cell mass in type 2 diabetes may be induced by different mechanisms, possibly acting in various combinations. There is now good evidence that cytokines, either produced by the beta-cells themselves in response to hyperglycemia [56] or by inﬂammatory cells, which localize at the periphery of the islets [57], can participate in the induction of beta-cell apoptosis. Adipokines, produced by fat cells, such as tumor necrosis factor-alpha (TNF-alpha), may also have deleterious effects on beta-cell survival [54]. Another apoptotic stimulus for beta-cells is the combination of elevated blood glucose and lipid concentrations, referred to as glucolipotoxicity [58]. Finally, more recent evidence

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indicates that ER stress, activating the Unfolded Protein Response (UPR), may cause beta-cell apoptosis [59,60]. Evidence has now been provided that each form of stress-induced apoptosis could be prevented or at least markedly reduced by GLP-1 or exendin-4 treatment. For instance, puriﬁed rat beta-cells exposed to cytokines were signiﬁcantly protected against apoptosis by treatment with exendin-4 [61], and a similar protection was also found in cytokine-treated INS-1E cells, an effect requiring an increase in Akt/PKB phosphorylation [62]. Protection against glucolipotoxicity was demonstrated in human islets exposed to high concentrations of glucose or palmitate, or a combination of both. In each condition, GLP-1 reduced the number of apoptotic cells by a mechanism involving Akt/PKB and nuclear factor-kappaB (NF-kappaB) activation [63]. More recently, it was shown that exendin-4 accelerates the recovery from translation inhibition caused by ER stress. This action of exendin-4 may ensure a fast return of insulin biosynthesis to normal rates as well as a stimulation of insulin secretion following UPR activation [64]. Consistent with in vitro data, administration of GLP-1 or exendin-4 to 8-week-old db/db mice or Zucker diabetic rats markedly decreased beta-cell apoptosis in pancreatic islets [53,65]. Studies of the signaling pathways mediating the antiapoptotic actions of GLP-1 support a role for PI3K, Akt/PKB, cAMP, and MAPK, in part via decreased levels of caspase-3 and increased levels of Bcl-2 and Bcl-XL [53,65,66]. The importance of the cAMP pathway has been studied in two animal models. Firstly, expression of a dominant-negative form of CREB in beta-cells results in a marked decrease in beta-cell mass and function [67] and causes a downregulation of the adaptor protein IRS-2. As IRS-2 is involved in the adaptation of beta-cells to metabolic demands [68], these data indicate a molecular link between the cAMP and IRS-2 signaling pathways in the control of beta-cell mass [69]. Secondly, transgenic mice expressing the negative regulator of CREB, inducible cAMP early repressor (ICER), in beta-cells show reduced beta-cell mass, hyperglycemia, and low plasma insulin levels. Interestingly, analysis of beta-cell proliferation and mass suggests that overexpression of ICER results in reduced proliferation rather than increased apoptosis [70]. Furthermore, recent evidence shows the importance of PDX-1 in the pleiotropic actions of GLP-1. Mice with a beta-cell speciﬁc inactivation of the Pdx1 gene exhibit a complete loss of the proliferative and anti-apoptotic effects of exendin-4 [71]. GLP-1 may use either IRS-2 or its downstream effectors PI3K or Akt/PKB [67,69] to upregulate PDX-1 expression [48]. This could be achieved by Akt-mediated phosphorylation of Foxo1 promoting its nuclear exclusion [72]. Moreover, GLP-1 increases forkhead box protein A2 (Foxa2) expression in association with decreased Foxo1 binding to the Foxa2 promoter [72]. This could prevent Foxo1 repressor effect on Foxa2-dependent

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stimulation of the PDX-1 promoter [73], resulting in activation of PDX-1 gene transcription, and expression of beta-cell genes as well as proliferative and survival factors.

Physiological Role of the Gluco-Incretin Hormones Assessed in Receptor Knockout Mice Most of the studies performed to evaluate the in vivo role of GIP or GLP-1 used pharmacological doses of these hormones or relied on inhibition of dipeptidylpeptidase IV, the ubiquitous enzyme that degrades these hormones. To evaluate the real physiological role of these hormones, mice with genetic inactivation of their receptors have been useful. Mice with inactivation of the GLP-1 receptor [74] were glucose intolerant following oral glucose administration and this was linked to a defect in insulin secretion, demonstrating the role of this hormone and its beta-cell receptor in the glycemic control during the absorptive phase. However, tolerance to intraperitoneal glucose administration was only mildly impaired. Thus, absence of this receptor did not lead to major dysfunction of glucose homeostasis. Inactivation of the GIP receptor in mice led to impaired tolerance to oral but not to intraperitoneal glucose administration [75]. Whereas high-fat feeding for 3 weeks induced a marked increase in the glycemic excursions following meal absorption, the absence of the GIP receptor actually prevented obesity development. The protective effect against obesity development of GIP receptor deletion was further shown by transferring the GIP receptor null alleles onto the ob/ob background. This markedly reduced body weight gain of the ob/ob mice, probably as a result of a reduced expression of the adipocyte genes controlling triglyceride deposition [76]. To evaluate whether simultaneous deletion of both the GLP-1 and GIP receptors would lead to more severe glucose intolerance, knockout mice for each receptor have been generated [77,78]. The double knockout mice had the same body weight and fasted or fed glycemia as wild-type or single receptor knockout mice. However, oral glucose administration revealed that the intolerance to glucose was additive with inactivation of both receptors and was correlated with a proportionate reduction in insulin secretion. Absence of both receptors led to intolerance to intraperitoneal glucose administration in female but not in male mice suggesting a possible defect in insulin secretion when both receptors are inactivated. This was tested in isolated islet perifusion studies which showed that glucose-stimulated insulin secretion was reduced by ∼50% whereas secretion from islets of mice with single receptor deletion was normal. The reduced secretory response in double receptor knockout islets could not be explained by a reduction in islet

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number, size or insulin content. Thus, absence of both gluco-incretin receptors led to a beta-cell autonomous secretion defect, which was found to be due to a defect distal to plasma membrane depolarization [77,78]. The secretion defect was not caused by a lower cAMP concentration in mutant islets. However, following forskolin treatment, which very strongly increases intracellular cAMP levels, the difference in secretion between control and mutant islets was abrogated. One possible explanation for this difference in secretion in the presence of different cAMP levels is that relatively low cAMP concentrations are sufﬁcient to induce the phosphorylation of speciﬁc PKA targets, since the regulatory subunit of PKA has a Kd for cAMP of ∼50 nM whereas Epac2 binds cAMP with a Kd in the micromolar range. Thus, forskolin-induced cAMP production may recruit the Epac2 pathway which may be similarly effective in triggering insulin secretion in both control and mutant islets. By contrast, at lower cAMP levels, when only PKA is activated, the reduced secretory response may result from the differential expression of speciﬁc PKA target proteins involved in insulin secretion. These PKA targets are still not identiﬁed.

Regulation of Beta-Cell Mass in the Absence of Gluco-Incretin Receptors GLP-1 or the agonist exendin-4 has been reported to increase beta-cell mass, especially in diabetic mice or rats [47,49,50,79]. Furthermore, a number of studies have provided evidence that GLP-1 can stimulate the differentiation of beta-cell precursors [42–44,47,80,81]. Inactivation of the GLP-1 receptor does not lead to any reduction in beta-cell mass in the adult mouse nor to a reduction in insulin content. However, impaired recovery of beta-cell mass after partial pancreatectomy in GLP-1 receptor knockout mice suggests that this receptor may participate in the control of beta-cell mass regeneration, but this appears to depend on the genetic background of the mice studied [82]. Thus, GLP-1 is probably only one of several mechanisms by which betacell mass can be regulated.

Regulation of the Hepatoportal Sensor by the GLP-1 Receptor As GLP-1 is secreted directly into the hepatic portal circulation, the highest concentration of the biologically active GLP-1-(7–36)–NH2 is found in the portal vein, where a glucose sensor is located. This hepatoportal glucose sensor is activated by a glucose gradient established between the portal and peripheral blood. This sensor activates glucose storage in liver, inhibits

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counterregulation, inhibits feeding, and stimulates glucose uptake by muscle and brown fat [83,84]. This sensor is connected to afferent vagal nerves projecting to the brainstem and hypothalamus. The ﬁring rate of these hepatic vagal afferents is modulated by glucose [85,86] and by GLP-1 [87], but not by the other incretin hormone GIP [88]. Activation of the portal sensor by portal vein GLP-1 infusion has been reported to increase glucose-stimulated insulin secretion [89]. We also showed that the GLP-1 receptor was required for the function of this sensor [90]. Indeed, we showed that in GLP-1 receptor knockout mice receiving an intraportal glucose infusion glucose clearance and muscle glucose uptake were no longer stimulated. A similar inhibition of glucose clearance was observed when exendin-(9–39) was co-infused with glucose into the portal vein of control mice. Thus, the glucose competence of the hepatoportal sensor requires the presence of the GLP-1 receptor. This is in contrast to the pancreatic beta-cells where the glucose competence is preserved in the receptor knockout animals [91]. An interesting role of the hepatoportal glucose sensor is the regulation of the ﬁrst-phase insulin secretion when glucose is injected intraperitoneally (i.p.). For instance, i.p. injection of glucose in GLP-1 receptor knockout mice fails to induce the ﬁrst phase of insulin secretion. Absence of ﬁrst-phase secretion was also observed in control mice receiving intraportal vein exendin(9–39), and in mice in which the hepatoportal sensor was inactivated by inactivation of the GLUT2 gene [77]. First-phase secretion was however preserved in the GIP receptor knockout mice, consistent with the absence of this receptor from the hepatoportal vein sensor. Importantly, islets isolated from control, GLP-1 receptor, or GIP receptor knockout mice, displayed the exact same kinetics of insulin secretion with brisk ﬁrst-phase insulin secretion when the glucose concentration was raised. These data suggest that in vivo, the main mechanism controlling ﬁrst-phase insulin secretion may be indirect, i.e., through activation of glucose sensors located outside of the endocrine pancreas, in particular in the hepatoportal vein region. This is also consistent with the observation that vagal afferents play an important role in the control of insulin secretion by physiological GLP-1 levels [92].

Conclusion Gluco-incretin hormones, in particular GLP-1 because of its potential use in the treatment of type 2 diabetes, have received considerable attention over the past 20 years. The effect of these hormones on the differentiation, proliferation, and function of pancreatic beta-cells is of considerable interest. However, the detailed molecular events activated following binding of these hormones to their cognate receptors, and which control beta-cell biology, are

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still far from being elucidated. Uncovering the complex signaling networks activated by these receptors and the transcription programs they control may lead to the identiﬁcation of potentially valuable novel targets that can be exploited for the development of new therapies for type 2 diabetes, but possibly also to increase beta-cell mass in type 1 diabetic patients. Finally, and importantly, studies with gene knockout mice have revealed that the control of insulin secretion in the intact animal may depend on GLP-1 action on other primary cellular targets, such as those forming the hepatoportal vein glucose sensors. As evidence strongly implicates this sensor in the control of ﬁrstphase insulin secretion, the disappearance of which is a sign of progression towards diabetes, deregulation of this sensor may be part of the very early events leading to abnormal insulin secretion in type 2 diabetes and which could be treated with GLP-1.

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27. Buteau J, Foisy S, Joly E, Prentki M (2003) Glucagon-like peptide 1 induces pancreatic beta-cell proliferation via transactivation of the epidermal growth factor receptor. Diabetes 52:124–132 28. Buteau J, Foisy S, Rhodes CJ, Carpenter L, Biden TJ, Prentki M (2001) Protein kinase Czeta activation mediates glucagon-like peptide-1-induced pancreatic beta-cell proliferation. Diabetes 50:2237–2243 29. Ebert R, Creutzfeldt W (1987) Gastrointestinal peptides and insulin secretion. Diabetes Metab Rev 3:1–26 30. Unger RH, Eisentraut AM (1969) Entero-insular axis. Arch Intern Med 123:261– 266 31. Holst JJ, Orskov C, Vagn Nielsen O, Schwartz TW (1987) Truncated glucagon-like peptide 1, an insulin-releasing hormone from the distal gut. FEBS Lett 211:169–174 32. Gromada J, Bokvist K, Ding WG, Holst JJ, Nielsen JH, Rorsman P (1998) Glucagon -like peptide 1 (7–36) amide stimulates exocytosis in human pancreatic beta-cells by both proximal and distal regulatory steps in stimulus-secretion coupling. Diabetes 47:57–65 33. Porksen N, Grofte B, Nyholm B, Holst JJ, Pincus SM, Veldhuis JD, Schmitz O, Butler PC (1998) Glucagon-like peptide 1 increases mass but not frequency or orderliness of pulsatile insulin secretion. Diabetes 47:45–49 34. Thorens B, Dériaz N, Bosco D, DeVos A, Pipeleers D, Schuit F, Meda P, Porret A (1996) Protein kinase A dependent phosphorylation of GLUT2 in pancreatic {beta} cells. J Biol Chem 271:8075–8081 35. Béguin P, Nagashima K, Nishimura M, Gonoi T, Seino S (1999) PKA-mediated phosphorylation of the human KATP channel: separate roles of Kir6.2 and SUR1 subunit phosphorylation. EMBO J 18:4722–4732 36. MacDonald PE, Wheeler MB (2003) Voltage-dependent K(+) channels in pancreatic beta cells: role, regulation and potential as therapeutic targets. Diabetologia 46:1046– 1062 37. MacDonald PE, Wang X, Xia F, El-kholy W, Targonsky ED, Tsushima RG, Wheeler MB (2003) Antagonism of rat beta-cell voltage-dependent K+ currents by exendin 4 requires dual activation of the cAMP/protein kinase A and phosphatidylinositol 3kinase signaling pathways. J Biol Chem 278:52446–52453 38. Ämmälä C, Ashcroft FM, Rorsman P (1993) Calcium-independent potentiation of insulin release by cyclic AMP in single {beta} cells. Nature 363:356–358 39. Ozaki N, Shibasaki T, Kashima Y, Miki T, Takahashi K, Ueno H, Sunaga Y, Yano M, Matsura Y, Iwanaga T, Takai Y, Seino S (2000) cAMP-GEFII is a direct target of cAMP in regulated exocytosis. Nat Cell Biol 2:805–811 40. Woodford TA, Correll LA, McKnight GS, Corbin JD (1989) Expression and characterization of mutant forms of the type I regulatory subunit of cAMP-dependent protein kinase. The effect of defective cAMP binding on holoenzyme activation. J Biol Chem 264:13321–13328 41. Holz GG, Leech CA, Heller RS, Castonguay M, Habener JF (1999) cAMP-dependent mobilization of intracellular Ca2+ stores by activation of ryanodine receptors in pancreatic β-cells. J Biol Chem 274:14147–14156 42. Zhou J, Wang X, Pineyro MA, Egan JM (1999) Glucagon-like peptide 1 and exendin-4 convert pancreatic AR42J cells into glucagon- and insulin-producing cells. Diabetes 48:2358–2366 43. Zhou J, Pineyro MA, Wang X, Doyle ME, Egan JM (2002) Exendin-4 differentiation of a human pancreatic duct cell line into endocrine cells: involvement of PDX-1 and HNF3beta transcription factors. J Cell Physiol 192:304–314

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17. Incretins and Regulation of Insulin Secretion Michael A. Nauck* and Juris J. Meier†

Summary. The incretin effect is the phenomenon whereby oral glucose elicits a greater insulin secretory response than an intravenous administration of glucose, even if the same glycemic proﬁle is obtained (“isoglycemia”) or even exceeded. The incretin effect mainly is the results of the secretion, from gut endocrine cells, of incretin hormones, which is stimulated by the ingestion and absorption of nutrients. The main incretin hormone is gastric inhibitory polypeptide (glucose-dependent insulinotropic polypeptide, GIP), produced in and secreted from upper intestinal K cells. A second incretin hormone, glucagon-like peptide-1 (GLP-1), is synthesized mainly in lower intestinal L cells. Both incretin hormones stimulate insulin secretion by interacting with speciﬁc receptors on endocrine pancreatic beta cells. This augmentation is prominent at high glucose concentrations, but stops at glucose concentrations slightly below fasting values. In patients with type 2 diabetes, the incretin effect is reduced and the reason is that GIP has lost most of its insulinotropic activity. GLP-1, on the other hand, has preserved activity, even in patients with type 2 diabetes. In addition to its insulinotropic activity, it also suppresses glucagon, retards gastric emptying, reduces appetite and food intake, and can inhibit beta-cell apoptosis and promote beta-cell regeneration and neogenesis. Therefore, these properties of GLP-1 can be exploited to treat type 2 diabetes, both in the form of incretin mimetics (GLP-1 receptor agonists) and DPP-4 inhibitors (preventing degradation and inactivation of incretin hormones by the proteolytic enzyme dipeptidyl peptidase-4). *Diabeteszentrum Bad Lauterberg, Kirchberg 21, D-37431 Bad Lauterberg, Germany † Medizinische Klinik I, St. Josef-Hospital, Klinikum der Ruhr-Universität Bochum, Gudrunstrasse 59, D-44791 Bochum, Germany

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The Incretin Effect in Healthy Subjects Ingestion of glucose leads to an approximately 5–10 fold stimulation of insulin secretion over the fasting, basal rate [1,2]. Part of this increment in insulin secretion is due to glucose being absorbed and leading to a rise in glycemia. However, this “direct” stimulation of insulin secretion through moderate elevations in circulating glucose concentrations only partially explains the insulin secretory response after glucose ingestion: if glucose is infused intravenously at rates that lead to a duplication of the glycemic excursion after oral glucose, a greatly reduced insulin and C-peptide response is observed [3–5]. This clearly indicates that other factors beyond a rise in glucose concentrations are important determinants of insulin responses after glucose ingestion [6,7]. Based on studies dating from 1906 showing the glucose lowering effects of gut mucosal extracts in diabetic patients [8], a possible explanation for differences in insulin secretory responses to oral and parenteral (e.g., intravenous) glucose was sought in the secretion and action of gut-derived hormones, the so called incretins [3,4,6,7]. Shortly after a method to measure insulin concentrations in plasma became available [9], a higher insulin response to oral as compared to intravenous glucose became evident [3,4], even in experiments that administered the same amount of glucose via both routes (thus leading to much higher circulating glucose concentrations when glucose was infused intravenously) [10,11]. Perley and Kipnis [12] introduced the method of “isoglycemic” intravenous glucose infusions to exactly compare the effects of oral and intravenous glucose under conditions, where the glycemic stimulus was the same. They described approximately three-fold higher insulin increments with oral than with intravenous glucose in healthy subjects. With advances in the methodology to assess insulin secretion (C-peptide, calculation of insulin secretion rates by deconvolution techniques) it became possible to quantify the incretin effect more deﬁnitively. Technically, the incretin effect is the difference in insulin secretory responses to oral and “isoglycemic” intravenous glucose (Fig. 1). Usually, it is expressed as the percentage of the (“total”) response after oral glucose (=100%). It makes a difference whether insulin, C-peptide, or insulin secretion rates are used to calculate the incretin effect [5,13]. The reason is that oral glucose leads to a change in insulin elimination form the circulation, most likely through a reduction in hepatic extraction [14], which lets more insulin appear in the general circulation. Since this is not the case with intravenous glucose (at least not to the same degree), incretin effects calculated from insulin responses are greater (60%–80% for a glucose load of 50 g) than incretin effects calculated from C-peptide or insulin secretion rates

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Fig. 1. Quantiﬁcation of the incretin effect in healthy subjects (left panels) and patients with type 2 diabetes. Glucose infusion rates (upper row), and concentrations of glucose (second row), insulin (third row), and C-peptide (bottom row) after oral glucose (red and blue symbols) and during “isoglycemic” glucose infusion (green and gray symbols) are shown. Asterisks indicate signiﬁcant differences between both routes of administering glucose. Adapted from Nauck et al. [15]

(approximately 40%–60%) [5,13,15]. The size of the incretin effects depends on the glucose load: it is small with minor oral glucose loads (like 25 g) and progressively increases with greater loads (e.g., 100 g), where it contributes approximately 80% to the responses in insulin and 60% to C-peptide and insulin secretion [5,10,11] (Fig. 2). Thus, the contribution of the incretin mechanism to insulin secretion after an oral glucose load is considerable. The incretin effect, therefore, constitutes a major mechanism for stimulating insulin secretion after nutrient ingestion, especially after glucose intake. A

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similar phenomenon exists regarding the comparison of protein ingestion and amino acid infusions [16]. It is not exactly known, what the incretin contribution is to insulin secretory responses after mixed meals. It may, however, be considered very likely that the insulin secretory responses following the ingestion of mixed meals greatly depends on contributions made by incretin-related mechanisms.

Incretin Hormones The incretin effect can in principle be explained by the secretion and insulinotropic action of hormones secreted in the course of nutrient assimilation (incretin hormones) [16–18] or by mechanisms involving the autonomous nervous system [19]. While there is evidence that nerve endings in the gut respond to the exposure to glucose and, possibly, other nutrients, and that endocrine pancreatic function is determined by sympathetic and parasympathetic input [20], pancreas transplantation does not modify the incretin effect [19,21]. Since it can be assumed that after heterotopic pancreas transplantation the functioning graft is denervated, this argues against a major role of neural mechanisms in mediating the incretin effect. The search for incretin hormones began with studies of gut-derived peptides known for other functions already, but also able to stimulate insulin secretion [17,22]. Such insulinotropic “side effects” (as a rule, at higher than physiological concentrations) were known, for example, for secretin, gastrin, cholecystokinin, vasoactive intestinal peptide (VIP), and others [6,7]. To be considered an incretin hormone, Creutzfeldt suggested that the following criteria had to be fulﬁlled: (1) release from gut endocrine cells after ingestion of nutrients, especially glucose; (2) stimulation of insulin release at typical postprandial concentrations; and (3) glucose dependence of the insulinotropic effects. Historically, the designation “incretin” is derived from secretin, which initially was a biologically active gut extract with no known chemical nature, and actions on both the exocrine and the endocrine secretions from the pancreas. Thus, theoretically, secretin was divided into “excretin” (stimulating exocrine secretion) and “incretin” (thought to act on endocrine pancreatic secretion). In 1971, gastric inhibitory polypeptide (GIP) was ﬁrst described by Brown et al. [23]. Its isolation was guided by a bioassay for gastric acid secretion. Shortly thereafter, the glucose-dependent insulinotropic action of GIP was described in the isolated perfused pancreas [24] and in vivo [18,25]. It became obvious that this insulinotropic activity represented the main biological action of GIP [26], which later was renamed “glucose-dependent insulinotropic polypeptide.” Thus, a ﬁrst incretin hormone was discovered [27]. The discovery of a second incretin hormone, glucagon-like peptide-1 (GLP-1), was a product of

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analyzing the proglucagon gene [28,29]. Two sequences not identical, but homologous to glucagon (“glucagon-like”) were described in 1983 [29]. At this stage, it was not known whether there would be a secreted peptide corresponding to the sequence of these two glucagon-like peptides (GLPs). Insulinotropic actions were among the ﬁrst effects looked for [30]. After deﬁning the true amino acid sequence as GLP-1 7–36(amide) [31,32], insulinotropic effects were deﬁned [33–35]. Later it became clear that GLP-1 is a multifunctional hormone with effects on other biological processes as well [36–39].

Gastric Inhibitory Polypeptide (Table 1) Gastric inhibitory polypeptide was the ﬁrst gut hormone that fulﬁlled the deﬁnition of an incretin hormone (see Creutzfeldt [6]): GIP is secreted in response to carbohydrate ingestion [40–42] and, in the concentrations reached after meals, is able to stimulate insulin secretion [18,43,44]. In fact, GIP augments the insulin secretory response to glucose. It has little ability to stimulate insulin secretion at normal fasting or lower glucose concentrations [24,26,35]. The higher the glucose concentration rises, the greater is its effect on insulin secretion.

Table 1. Comparison of the biological characteristics and actions of gastric inhibitory polypeptide (GIP) and glucagon-like peptide 1 (GLP-1) GIP References GLP-1 Biological characteristics Secretion Size Stimulus for secretion Plasma concentrations Fasting Postprandial Half life (intact forms) Physiological actions Insulin secretion Glucagon secretion Gastric emptying Gastric acid secretion Satiety/food intake Beta-cell apoptosis Beta-cell regeneration Lipid homeostasis

K cells (mainly duodenum/jejunum) 42 amino acids Fat/carbohydrates

L cells (mainly colon, ileum) 29 amino acids Fat/carbohydrates

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[27,29] [51,207]

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GLP-1 (7-36 amide) His Ala Glu Gly Thr Phe Thr Ser Asp Val Ser Ser Tyr Leu Glu Gly Gln Ala Ala Lys Glu Phe Ile Ala Trp Leu Val Lys Gly Arg -NH2

GIP (1-42 amide) Tyr Ala GluGly Thr Phe Ile Ser Asp Tyr Ser Ile Ala MetAsp Lys Ile His Gln GlnAsp PheVal Asn Trp Leu Leu Ala GlnLys Gly Lys LysAsnAspTrpLys His Asn Ile ThrGln -NH2

Site of proteolytic inactivation (DPP-4)

Fig. 3. Amino acid sequence of the incretin hormones gastric inhibitory polypeptide (GIP) and glucagon-like peptide-1 (GLP-1). Identical amino acids are shown in blue, amino acids unique to GLP-1 and GIP are shown in green and red, respectively

Gastric inhibitory polypeptide is produced in endocrine cells located as single cells within the mucosa of the upper small intestine (duodenum and upper jejunum), the so-called K-cells [45]. It is secreted after the ingestion of carbohydrates, especially glucose and galactose [42], triglycerides [46], and certain amino acids [47,48]. It is synthesized as a larger precursor peptide (proGIP) [49] and through post-translational processing becomes the 42 amino acid, C-terminally amidated incretin known as GIP [50] (amino acid sequence, see Fig. 3). Fasting plasma concentrations are 10–50 pmol/l, and soon after nutrient ingestion, concentrations rise to 80–400 pmol/l and remain elevated for several hours [51]. These numbers refer to nonspeciﬁc assays that do not differentiate between the intact GIP peptide and des-Tyr-Ala-GIP degraded by dipeptidyl peptidase-4 and represent “total” GIP. Assays detecting intact GIP more speciﬁcally read lower concentrations, representing 40%–70% of total GIP [50,52–54]. Gastric inhibitory polypeptide acts through speciﬁc receptors belonging to the 7-trans-membrane-domain family of receptors [55–58]. They are expressed in the endocrine pancreas, on adipocytes, and in the brain. Apart from stimulating insulin secretion, GIP can, under certain circumstances, raise glucagon secretion [26,59,60]. In addition, a role for GIP in promoting triglyceride storage in adipose tissue (and, hence, weight gain) has been suggested [61,62].

Glucagon-Like Peptide-1 (Table 1) Glucagon-like peptide-1 is one product of L cells most predominantly situated in the lower small and large intestines [32,63,64]. The same cells secrete enteroglucagon (glicentin), GLP-2, and peptide tyrosine tyrosine (PYY) [65].

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Glucagon-like peptide-1 is one product of post-translational processing of proglucagon in L cells [31,32,66]. It corresponds to amino acids 78–107 of the proglucagon sequence (amino acid sequence, see Fig. 3). Regarding the carboxy-terminus there are two forms of GLP-1: The amidated form GLP1(7–36) amide [32,67] and the glycine-extended form GLP-1(7–37) [31,68]. The amidated form appears to be more predominant [67]. No differences have been described regarding biological effects or pharmacokinetic behavior [69]. Plasma concentrations for GLP-1 are lower than described for GIP: in the fasting state concentrations of 2–15 pmol/l have been described. After nutrient ingestion, these levels rise to 15–45 pmol/l [67,70,71]. These concentrations, again, represent “total GLP-1” concentrations. In the case of GLP-1, intact, biologically active concentrations have been measured as a rather low proportion of total concentrations, in the range of 10%–20% [72]. This indicates that GLP-1 is a high afﬁnity substrate for the ubiquitous exopeptidase dipeptidyl peptidase-4 (DPP-4) [73]. Glucagon-like peptide-1 receptors are highly speciﬁc and also belong to the 7-trans-membrane-domain family of receptors [74]. They are expressed in the endocrine pancreas, in the autonomous nervous system, in the brain, and elsewhere [75]. This indicates that GLP-1 does not only affect insulin secretion [37,39,76–78]. It is more than an incretin hormone [79], which makes the deﬁnition of its true physiological role more complex [80,81].

Other Incretin Candidates There is an ongoing debate as to the existence of additional incretin hormones in humans. This assumption has been based on earlier experiments (prior to the discovery of GLP-1) showing that intestinal extracts, in which GIP had been neutralized by speciﬁc immunoblockade, still exerted a signiﬁcant incretin activity [82]. However, subsequent studies demonstrated that most of this activity could be ascribed to the actions of GLP-1 [35,83]. Nevertheless, the possibility remains that other hormones besides GIP and GLP-1 are involved in the physiological incretin effect in humans. Cholecystokinin One candidate hormone for an incretin role was cholecystokinin (CCK). Indeed, experiments in isolated perfused pancreas preparations as well as in vivo in mice and rats studies glucose-dependent insulinotropic properties for CCK [84–86], even though at more physiological plasma levels these effects were rather modest [87]. Furthermore, reduced circulating plasma concentrations of CCK in patients with type 2 diabetes pointed to a role for CCK in the pathogenesis of impaired insulin secretion in these patients [88]. However, some of the earlier studies need to be regarded with caution since crude

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preparations of CCK eventually also contained GIP, thereby not allowing for a clear distinction between the actions of these two gut hormones [89]. Moreover, there is a large heterogeneity regarding the insulinotropic effects of CCK in different species, and its efﬁcacy in humans appears to be particularly small [90]. Thus, in healthy humans, endogenously secreted CCK could not be linked to an increment in insulin release, and exogenous CCK failed to enhance insulin secretion at typical physiological postprandial plasma concentrations [91]. Also, the insulinotropic effect of GIP could not be augmented by the co-administration of CCK [44]. Finally, even though CCK is secreted in response to meal ingestion, glucose in particular appears to be a weak stimulus [92], and administration of the CCK antagonist loxiglumide did not lead to a substantial reduction in the incretin effect [93–95]. Therefore, the incretin activity of CCK in humans appears to be negligible. Nevertheless, this hormone contributes to postprandial glucose regulation via inhibiting gastric emptying and consequently delayed entry of nutrients into the circulation [96]. Gastrin Gastrin is another gastrointestinal hormone suggested to possess incretin activity. Circulating gastrin levels rise by ∼2-fold after oral glucose or meal ingestion [97,98], and exogenous administration of gastrin was shown to elicit a glucose-dependent increase in insulin concentrations in some [97,99–101], but not all studies [102–105]. However, dose–response studies revealed that augmentation of insulin release was only achieved at supraphysiological plasma levels [97], and the modest increments in circulating gastrin concentrations measured after oral glucose ingestion in man appeared to be of insufﬁcient magnitude to have an impact on insulin secretion [105]. Thus, at physiological concentrations gastrin seems to have little, if any, importance for glucose control in humans, and therefore cannot be classiﬁed an incretin hormone. Secretin Like CCK, secretin is secreted in response to nutrient ingestion, and glucose in particular appears to be a weak stimulus [106]. Insulin secretion can be augmented in a glucose-dependent manner by secretin in rodents as well as in humans, but the magnitude of this effect is rather small [100,107–110]. Overall, the weak stimulation of secretin release after oral glucose and its modest insulinotropic effects at physiological plasma levels do not qualify this peptide as a typical incretin hormone. PACAP and VIP The neuropeptides PACAP (pituitary adenylate cyclase-activating polypeptide) and VIP (vasoactive intestinal polypeptide) stimulate insulin secretion in a glucose-dependent manner [111], and antagonizing the PACAP receptor in mice leads to disturbances in postprandial insulin secretion [112]. However,

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both peptides primarily act as neurotransmitters rather than as typical hormones, and therefore cannot be considered classical incretin hormones. GLP-2 Glucagon-like peptide 2 (GLP-2), like GLP-1, arises as a product of posttranslational cleavage of the proglucagon gene product in intestinal L cells, and hence is secreted at equimolar amounts with GLP-1 [29,63]. However, while the concurrent secretion of GLP-1 and GLP-2 would suggest similar physiological actions, GLP-2 has no effect on insulin secretion and rather stimulates pancreatic glucagon release [30,104]. Therefore, GLP-2, if anything, even counters the incretin actions of GIP and GLP-1.

Glucose Dependence of Insulinotropic Actions of Incretin Hormones A remarkable feature differentiating both incretin hormones from available insulin-releasing drugs like sulfonylureas is the fact that they augment the insulin secretory response to hyperglycemia, but do no stimulate insulin secretion in the absence of a permissive glucose concentration [18,24,26,35,68]. In fact, at very low glucose concentrations GIP and GLP-1 are not able to stimulate insulin secretion at all [113,114]. The mechanism of stimulating insulin secretion in both the cases of GIP and GLP-1 involves cyclic AMP. Gastric inhibitory polypeptide and GLP-1 receptors are coupled to stimulating G proteins and activate adenylate cyclase [115– 117]. cAMP acts via protein kinase A to have an inﬂuence on multiple processes related to insulin secretion, as described in detail by Gromada et al. [117,118] (Fig. 4). The post-receptor-binding events appear to be indistinguishable for GIP and GLP-1 in islets/beta cells from healthy animals or human subjects. Elevated cAMP level can have an inﬂuence on the closure of the ATPdependent potassium channel, voltage-gated calcium channels, and on the process of exocytosis. The fact that incretins alone (i.e., in the absence of an elevated glucose concentration) cannot close the ATP-dependent potassium channel (whereas sulfonylureas can) at low glucose concentrations, or when elevated glucose concentrations return to lower values, explains why their action on insulin secretion is so explicitly glucose-dependent. This is the reason for the fact that it has not been possible to elicit an inadequate burst of insulin secretion leading to hypoglycemia by administering an “overdose” of GIP or GLP-1 to animals of human subjects [113,119]. Only if GLP-1 is coadministered with large amounts of glucose, and if the administration of glucose is stopped rapidly, will this highly artiﬁcial situation lead to reactive hypoglycemia [120]. Glucose dependence of insulinotropic actions of GIP and GLP-1 has been shown in the isolated perfused rat pancreas [24,26,68] as well as in vivo [18,35,114]. Kreymann et al. described results of physiological-dose infusions

Incretins and Regulation of Insulin Secretion Glucose, amino acids GIP

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Fig. 4. Schematic diagram of pancreatic beta cell. Receptors, enzymes, substrates, receptors, and second messengers involved in the insulin secretory process are shown. Glucagon-like peptide-1 binds to its receptor; this increases cAMP and activates protein kinase A. Activated protein kinase A can inﬂuence insulin secretion by effects on (a) the ATPdependent potassium channel, (b) on voltage-gated calcium channels, (c) on calcium released from intracellular stores, and, mainly, (d) on the process of exocytosis. Glucagonlike peptide-1 cannot, however, by itself close the ATP-dependent potassium channel and initiate insulin secretion independent from elevations in glucose concentrations. Adapted from Gromada et al. [116–118,272]

of GIP and GLP-1 in healthy subjects [35]. When the incretin hormones were infused in the fasting state, little effect on insulin or reduction in glucose concentrations was observed. If, however, an infusion of glucose was started in order to elevate glucose concentrations into the postprandial range, insulin secretion was augmented several-fold over the values observed with placebo. When GLP-1 was infused at a pharmacological dose (leading to “therapeutic, antidiabetic” levels of GLP-1) in the course of a stepwise hypoglycemic clamp experiment, insulin secretion (calculated by deconvolution of C-peptide concentrations) was stimulated at basal plasma concentrations in healthy subjects [114]. The effect was lower at lower glucose concentrations and negligible at any glucose concentration below 70 mg/dl (Fig. 5). Similar results

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Fig. 5A,B. Glucose dependence of insulin secretion stimulated by GLP-1 in healthy subjects. Hyperinsulinemic, step-wise hypoglycemic clamp experiments were performed in the presence of a pharmacological infusion of GLP-1(7–36) amide (1.2 pmol/kg per minute). The glucose plateaus are indicated. C-peptide (A) was measured. Insulin secretion rates (B) were derived by deconvolution. Ratios between values determined with GLP-1 over those with placebo were calculated. Mean values and 95% conﬁdence intervals are depicted. Only above 3.7 mmol/l glucose was GLP-1 able to signiﬁcantly stimulate insulin secretion. Adapted from Nauck et al. [114]

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have been obtained with exendin-4, another GLP-1 receptor agonist, in healthy subjects [121], or with liraglutide, a GLP-1 analogue, in type 2diabetic patients [122]. This points to the fact, underscored by other ﬁndings, that the threshold for insulinotropic effects of GLP-1 or the glucosedependence of such effects follow a similar pattern in healthy subjects and type 2 diabetic patients [123].

Additional Antidiabetogenic Actions of Incretin Hormones In addition to its glucose-dependent actions on endocrine pancreatic secretion, GLP-1 modulates glycemia through gastrointestinal and central nervous mechanisms, as well as perhaps via direct actions in the peripheral tissues.

Gastric Emptying The rate of gastric emptying modulates postprandial glucose homeostasis through regulation of nutrient inﬂow into the circulation. Horowitz et al. estimated the contribution of gastric emptying to the overall variations in postprandial glycemia to be in the order of 35% [124]. Thus, any deceleration of gastric emptying will blunt the postprandial rise in glycemia, whereas rapid emptying, particularly of high-glucose containing meals, can result in a rapid increase in circulating glucose levels, thereby causing exaggerated insulin responses (as is the case in patients with gastric dumping syndromes following resection of the stomach [125]). Amongst the substances known to decelerate gastric emptying, GLP-1 is probably one of the most potent ones. In studies with the intravenous infusion of different GLP-1 doses in humans, emptying of both liquid and solid meals was markedly delayed by the incretin hormone [79,126,127]. These effects were detectable at near-physiological GLP-1 plasma levels (∼50 pmol/l) [79,127], suggesting a physiological role for endogenous GLP-1 as an enterogastrone, i.e., as a gut hormone that acts to delay gastrointestinal transit time after meal ingestion [80]. At higher plasma concentrations (∼150 pmol/l), gastric motility was almost completely arrested, with ∼80% of the initial content still retained in the stomach four hours after the ingestion of a solid test meal [127]. In addition, GLP-1 also acts to inhibit gastric acid secretion [126,128]. The importance of the gastric emptying effects of GLP-1 is illustrated by the fact that postprandial insulin responses are reduced by GLP-1 administration, despite its insulinotropic actions [79,127,129]. This apparent paradox can be explained by the delayed entry of glucose intro the blood stream, whereby the direct nutrient stimulation of

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insulin secretion is prevented. In contrast, when the effects of GLP-1 on the stomach are countered (e.g., by the concomitant administration of the prokinetic drug erythromycin), the postprandial insulin responses are augmented by GLP-1 [129]. The mechanisms by which GLP-1 controls gastric emptying include both an increase in the pyloric tone as well as an inhibition of antroduodenal motility [130–132]. Most likely, these effects are mediated by an inhibition of efferent vagal activity, either via GLP-1 binding to central nervous receptors or to gastric GLP-1 receptors activating vagal projection ﬁbers to the hypothalamus [133]. The importance of the vagal nervous system for the GLP-1 effects on the stomach becomes evident from experiments in vagotomized rats [134] and humans [135] as well as from the inhibition of pancreatic polypeptide secretion by exogenous GLP-1 [129]. In contrast to GLP-1, GIP does not seem to have a signiﬁcant effect on gastric emptying or gastric acid secretion in humans [136,137].

Satiety and Food Intake Glucagon-like peptide-1 also exerts important effects on the regulation of satiety and food intake [133]. Along these lines, studies by Schick et al. and by Turton et al. demonstrated that the intracerebroventricular administration of GLP-1 reduces food intake in rats [138,139], and these effects could be antagonized by the co-administration of the GLP-1 receptor antagonist exendin(9–39) [139]. Consistent with these acute effects, long-term administration of the GLP-1 receptor agonist exendin-4 elicited a signiﬁcant reduction in body weight in rats [140]. In humans, Flint and colleagues as well as Gutzwiller and colleagues showed that intravenous administration of GLP-1 reduces appetite sensations and lowers food intake [141,142]. These results gave rise to suggesting GLP-1 as a potential anti-obesity drug [133]. Indeed, continuous subcutaneous infusion of GLP-1 in obese patients with type 2 diabetes over 6 weeks not only reduced the levels of glycemia, but also led to a mean weight reduction of ∼2 kg compared to placebo administration [143]. Whether the induction of satiety by GLP-1 is primarily based on a direct interaction of GLP-1 with its central nervous receptors, or whether this is secondary to the delay in gastric emptying is a current matter of debate [133]. Gastric distension is well known to result in a loss of appetite [144]. Central nervous GLP-1 receptors are also accessible to peripheral GLP-1 [145], and in rodents direct activation of such central GLP-1 receptors signiﬁcantly reduces the caloric intake [139,146]. Therefore, most likely both mechanisms contribute to the anorexic effects of GLP-1. Long-term studies using the GLP-1 derivative exenatide in patients with type 2 diabetes over more than 100 weeks have conﬁrmed its weight-reducing effect [147–150]. It seems noteworthy that in contrast to other anti-obesity

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drugs, such as rimonabant [151], the GLP-1 effects on body weight do not seem to level off after a certain period of treatment, but continue even during its long-term administration [150]. Since any reduction in body weight is associated with improved insulin sensitivity, the GLP-1 effects on body weight certainly contribute to its antidiabetogenic effects over longer periods of treatment. The effects of GIP on the regulation of appetite and food intake have not been studied so far.

Endogenous Glucose Production and Glucose Disposal Whether GLP-1 also exerts direct effects on glucose disposal has been widely debated [152]. Glucagon-like peptide-1 receptor binding has been reported in muscle and adipose tissue in some [153–155], but not all studies [156,157], and in isolated myotubes as well as in adipocytes direct insulinlike effects of GLP-1 have been found [158,159]. It is, however, unclear whether these effects play a role at physiological GLP-1 plasma levels in vivo. D’Alessio et al. reported increased glucose-disposal following GLP-1 administration in healthy humans using the minimal model approach [160,161]. In addition, Prigeon et al. showed that GLP-1 elicited a small, but signiﬁcant suppression of hepatic glucose production using a pancreatic clamp method [162], whereby plasma levels of insulin and glucagon can be matched by means of a somatostatin infusion. However, other investigators failed to conﬁrm these effects in healthy human subjects or patients with diabetes [163–165]. The reasons for these discrepancies are not entirely clear, but may be due to methodological issues. Measuring glucose disposal after intravenous glucose infusion following GLP-1 or placebo administration is complicated by the differences in insulin and glucagon levels between the tests [152]. The pancreatic clamp technique on the other hand allows for control of these factors, but might interfere with glucose homeostasis due to the metabolic effects of somatostatin itself. Therefore, it is possible that minor differences in glucose disposal are obscured by the complexity of this experimental system. One hypothesis that has been put forward is that the putative effects of GLP-1 on glucose disposal are mediated through its primary metabolite GLP1(9–36) amide [166–168], the circulating plasma concentrations of which markedly exceed those of intact GLP-1(7–36) amide [169]. In support of this thesis, Deacon et al. showed that intravenous administration of GLP-1(9–36) amide enhanced glucose elimination in pigs [166], and Nikolaidis et al. suggested that the positive inotropic effects of GLP-1 are also mediated through the GLP-1 metabolite [167]. When administered into healthy human subjects, no effects on the disposal of intravenously injected glucose were found by Vahl et al. [170]. In contrast, experiments from our group revealed a small,

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but signiﬁcant decrement in the postprandial rise of glucose concentrations during the infusion of GLP-1(9–36) amide, and these effects were independent of changes in insulin and glucagon secretion or gastric emptying [168]. Therefore, there is some evidence to suggest that GLP-1 does exert insulinmimetic effects on hepatic glucose production and peripheral glucose disposal, and that these effects are at least in part mediated through the metabolite GLP-1(9–36) amide. However, the overall importance of these effects at physiological plasma levels appears negligible. Aside from these potential direct actions on glucose disposal and endogenous glucose release, a signiﬁcant improvement of insulin sensitivity has been found after the chronic administration of GLP-1 [143]. However, when insulin sensitivity was directly measured during an acute GLP-1 infusion, no effects were found [171]. Thus, the improvements in insulin sensitivity observed during chronic GLP-1 administration are most likely secondary to the overall improvement of glycemic control, independent of any direct GLP-1 effect on the insulin-responsive tissues. For GIP, no direct effects on glucose production, glucose disposal, or insulin sensitivity have been described.

Beta-Cell Apoptosis and Regeneration Both type 1 and type 2 diabetes develop as a consequence of signiﬁcant betacell loss (∼99% beta-cell deﬁcit in type 1 diabetes, ∼60% deﬁcit in type 2 diabetes), presumably due to increased beta-cell apoptosis [172,173]. Therapeutic strategies aiming to enhance beta-cell regeneration and to inhibit beta-cell apoptosis might therefore prevent or at least delay the onset of diabetes in individuals at high risk [174]. A number of peptide hormones, including gastrin, GLP-2, IGF-1, growth hormone, and insulin, are known to modulate beta-cell proliferation and apoptosis [175–177]. Beta-cell proliferative effects of GLP-1 were ﬁrst reported in a beta-cell line (INS-1 cells), where it increased thymidine incorporation via induction of phosphatidylinositol 3-kinase, protein kinase Czeta, activation of PDX1 gene expression, and inactivation of the forkhead transcription factor Foxo1 [178,179]. Glucagon-like peptide-1 was also shown to increase beta-cell replication as well as putative islet neogenesis from ductal precursor cells in different rodent models [180–184]. Furthermore, under culture conditions exocrine pancreatic cells were converted into insulin-secreting cells by GLP-1 treatment [185,186], but it is questionable whether this pathway plays a role in vivo [187]. Glucagon-like peptide-1 was also reported to inhibit [184,188,189], including beta-cell apoptosis isolated human islets [188]. Like GLP-1, GIP inhibits beta-cell apoptosis and stimulates beta-cell proliferation through a phosphatidylinositol 3-kinase (PI3K)/protein kinase B (PKB) dependent pathway and inactivation of Foxo1 in INS-1 cells as well as

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isolated mouse islets [190,191]. Consistent with the actions of GIP and GLP-1, beta-cell preservation has also been shown in streptozotocin-induced diabetic rats after treatment with a DPP-4 inhibitor [192,193]. Since any interference with cell cycle regulation theoretically bears the risk of inducing cell autonomy, there has been some concern that the beta-cell proliferative effects of GIP and GLP-1 might result in tumor formation. However, recent studies provided evidence that neither acute, nor chronic stimulation of the GLP-1 receptor stimulated the growth or survival of human pancreatic cancer cells [194]. While the effects of GIP and GLP-1 on beta-cell turnover have been convincingly demonstrated in a number of in vitro and animal studies, the question remains as to whether this will indeed translate into actual regeneration or preservation of beta-cell mass in patients with diabetes. In fact, current evidence suggests that in humans the capacity of the islet beta cells to regenerate is rather low compared to that in mice or rats [172,174]. Furthermore, it is as yet impossible to directly measure such changes in beta-cell mass or turnover in humans, since the pancreas is inaccessible for repeated biopsy sampling, and functional assessment of insulin secretion may only partly relate to the actual beta-cell mass [195]. Service and colleagues have recently suggested that the excessive endogenous secretion of GIP and GLP-1 following gastric bypass surgery might lead to islet hyperplasia and nesidioblastosis resulting in the occurrence of hyperinsulinemic hypoglycemia [196]. However, on further analysis, these symptoms were linked to a functional hyperactivity of the beta cells rather than increased beta-cell mass or turnover [197]. Therefore, at present it cannot be answered with certainty whether GIP and GLP-1 have an impact on beta-cell regeneration in humans.

The Physiological Roles of GIP and GLP-1 Gastric inhibitory polypeptide is clearly an incretin hormone in animals [198] and healthy human subjects [44,83], with some additional actions, that probably are secondary in importance to its incretin role. Gastric inhibitory polypeptide has no effects on the motility of the stomach or gut [199]. This is different than GLP-1. The retardation of gastric emptying determines changes in the time course and activity of events that occur secondarily to delivery of nutrients into the duodenum [79]. Therefore, exposure to GLP-1 means delayed and slowed absorption of the carbohydrate component of a meal, smaller rises in glycemia, with less stimulation of insulin secretion, despite potential direct actions on the endocrine pancreas that would augment insulin responses. The net effect of exposure to GLP-1 may, therefore, well be a reduction rather than an augmentation of insulin secretory responses after a meal [79]. This is not easily compatible with a simple role as an incretin

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hormone [80]. As far as insulinotropic effects of are concerned, GIP and GLP1 act in an additive manner [83,200]. Given the differences in plasma concentrations (higher for GIP than for GLP-1) and the components of the action proﬁle of GLP-1 leading to a reduction rather than stimulation of insulin secretory responses, it is very likely that GIP is the major incretin in human physiology, with GLP-1 making a contribution that is probably smaller than that of GIP.

Incretin Effect in Type 2 Diabetes When the incretin effect was quantiﬁed in patients with type 2 diabetes, it was found to be reduced [15] (Fig. 5). This was evident in the seminal studies of Perley and Kipnis [12] in which only insulin responses were evaluated. When insulin and C-peptide responses were assessed after oral glucose and during “isoglycemic” intravenous glucose infusions, the incretin effect was found to be reduced or absent in patients with type 2 diabetes [15]. While there was a residual incretin effect if only insulin responses were examined, C-peptide concentrations (more precisely related to insulin secretion [201]) indicated no signiﬁcant difference in the ability of oral and intravenous glucose to stimulate insulin secretion in these type 2 diabetic patients, in contrast to age-matched control subjects with normal oral glucose tolerance [15]. The ﬁrst question arising from the reduced incretin effect in type 2 diabetes is that of the mechanism explaining the loss of the incretin effect. In principle, it may be related to defects in the secretion or insulinotropic action of GIP and/or GLP-1. The second relates the reduced incretin effect to the well-known abnormalities in insulin secretion in type 2 diabetic patients especially in the postprandial state [1,2]. If insulin secretion following nutrient ingestion in healthy subjects relies heavily on incretin-related mechanisms, which contribute approximately half of the overall insulin secretory response [5,10,13], should the absence of activity have important physiological consequences? We suggest that the defective pattern of insulin secretion known for type 2 diabetic patients, i.e., the initial delay and reduced responsiveness following a meal, is related to the lack of stimulation through incretin-related mechanisms.

Secretion of Incretin Hormones in Type 2 Diabetes The question in relation to the secretion of incretin hormones is whether absent or reduced responses after meals might help to explain the absent or reduced incretin effect. There is no lack of GIP secretion in patients with

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type 2 diabetes. Often, some over-secretion of GIP has been described when meal-related GIP responses have been compared between healthy subjects and type 2 diabetic patients [202]. Other studies have found no difference [203,204]. Obese subjects tend to have higher responses independent of abnormalities in glucose metabolism [205]. Studies involving larger groups suggest that there is heterogeneity in type 2 diabetic patients regarding the secretion of GIP. While there are subgroups with GIP hypersecretion, the majority of type 2 diabetic patients have normal GIP responses, and there may even be some with subnormal responses [7,206]. Overall, these subtle abnormalities do not explain the reduced incretin effect in patients with type 2 diabetes. The situation is somewhat different with regard to GLP-1. Initially, studies of small numbers of subjects reported contradicting results [207,208]. A large study involving subjects with normal and impaired glucose tolerance as well as type 2 diabetic patients has been presented by Toft-Nielsen et al. [203]. They describe a signiﬁcant reduction in “total” GLP-1 responses for type 2 diabetic patients and a similar trend for subjects with impaired glucose tolerance. The difference in GLP-1 plasma levels was more pronounced during the second and third hour after meal ingestion and amounted to approximately 25% of the integrated incremental GLP-1 responses (Fig. 6). Vilsbøll et al. supplemented these ﬁndings with a further analysis of “intact, biologically active” GLP-1, conﬁrming reduced GLP-1 responses in type 2 diabetes, both for “total” (the amount secreted) and “intact” GLP-1 [204].

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Fig. 6. Secretion of GLP-1 in patients with type 2 diabetes (red), subjects with impaired oral glucose tolerance (IGT, blue) and in healthy controls (green). The secretion of GLP-1 is signiﬁcantly diminished in patients with type 2 diabetes, whereas results in IGT subjects are intermediate. Adapted from Toft-Nielsen et al. [203]

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Actions of Incretin Hormones in Type 2 Diabetes Several studies have examined insulinotropic actions of GIP in patients with type 2 diabetes and found greatly reduced insulin-stimulating effectiveness [208–214] (Fig. 7) thus providing an excellent explanation for the reduced incretin effect in these patients. Recently, Vilsbøll et al. described the lack of insulinotropic activity of GIP even at extraordinarily high doses/concentrations, when infused continuously. Glucagon-like peptide-1 administered in the same manner was exquisitely active. On the other hand, when GIP was administered as a bolus injection, it retained a much higher relative activity in type 2 diabetic patients compared to healthy subjects [214]. Glucagon-like peptide-1, on the other hand, is much more active as an insulinotropic agent in patients with type 2 diabetes (Fig. 7). When GLP-1 was infused under hyperglycemic clamp conditions, only a minor (nonsigniﬁcant) difference in insulinotropic activity was noted when patients with type 2 diabetes were compared with age- and weight-matched control subjects with normal glucose tolerance [208]. In the same study, the action of GIP was heavily impaired in type 2 diabetic patients. Since both a physiological and a pharmacological dose of GLP-1 were tested in this study, one must conclude that GLP-1, in type 2 diabetic patients, has major insulinotropic activity only at pharmacological doses (1.2 pmol/kg per minute) and concentrations (exceeding postprandial concentrations by three-fold) [208]. This is consistent with the ability of pharmacological doses of GLP-1 to lower, and even normalize glucose concentrations in type 2 diabetic patients [123,215]. Regarding the relative insulinotropic potency of GLP-1 in patients with type 2 diabetes, Kjems et al. performed a detailed study investigating the dosedependence and the relationship between insulin secretion rates and glucose concentrations. They found that 0.5 pmol/kg per minute GLP-1 restored glucose-responsiveness in type 2 diabetic patients to normal [216]. This means that, with exposure to exogenous GLP-1, insulin secretion responds to an incremental infusion of glucose in the same way as is seen in healthy controls in the absence of exogenous GLP-1. Nevertheless, when glucose-responsiveness is compared over a range of different doses of exogenous GLP-1, it is approximately one-quarter to one-third in patients with type 2 diabetes relative to healthy controls [216]. This indicates that the insulinotropic action of GLP-1 in type 2 diabetic patients is not normal, but impaired. The relative impairment in the insulinotropic activity of GLP-1 compared to GIP in type 2 diabetic patients is less pronounced. It is not known whether the difference in the insulinotropic activity between GIP and GLP-1 is due to the relatively well-preserved activity of GLP-1 (compared to other insulin secretagogues) or to a comparably impaired activity of

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Type 2 diabetic patients

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AUCC-peptide [nmol.l-1.min]

AUCC-peptide [nmol .l-1.min]

Incretins and Regulation of Insulin Secretion

Fig. 7. Impaired insulinotropic effectiveness of GIP in patients with type 2 diabetes (upper panel) in comparison to healthy subjects. Area under the curve (AUC) C-peptide response to the infusion of physiological replacement doses and pharmacological doses of GIP during a hyperglycemic clamp experiment (8.5 mmol/l) are shown. For comparison, the response to physiological replacement doses and pharmacological doses GLP-1(7–36) amide are also depicted. Type 2 diabetic patients responded considerably less than healthy subjects to exogenous GIP, but responses to GLP-1 were similar. Adapted from Nauck et al. [208]

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GIP. One can only speculate with regard to mechanism(s). The ZDF rat model of type 2 diabetes is characterized by reduced expression of GIP receptors in the endocrine pancreas [217]. However, the relatively well preserved activity of GIP after bolus injection [214] argues against deﬁcient GIP receptors as a major cause of the impaired response to GIP, but does not exclude alterations in their functional status. There could, for example, be changes favoring desensitization [218] upon prolonged exposure. However, repeated bolus injections do not lead to impaired insulin secretory responses with the second exposure to GIP in patients with type 2 diabetes [219]. Another riddle concerns the question as to why the insulinotropic effects of GLP-1 and GIP are affected so differently in type 2 diabetes, although the mechanism of action of GLP-1 and GIP are so similar. Thus, it is difﬁcult to identify a step in the post-receptor signaling pathways, where the difference may be located. The reduced insulinotropic effectiveness of GIP is not restricted to type 2 diabetes, but is observed in all other forms of diabetes examined: latent autoimmune diabetes in adults (LADA) with preserved insulin secretory capacity; diabetes secondary to chronic pancreatitis; and MODY-2 diabetes [220]. This indicates that the loss of insulinotropic activity of GIP may not be a primary, but secondary to the diabetic (hyperglycemic) state. Changes in GIP receptor number and activity have been described in islets exposed to hyperglycemia [217,221]. The loss of insulinotropic GIP activity needs to be examined during the natural history of the development of type diabetes to determine if it is reversible.

Secretion and Action of Incretin Hormones in Prediabetic States The marked reduction of the incretin effect in patients with overt diabetes gives rise to the question whether the defects in the entero-insular axis in patients with type 2 diabetes might be due to a primary, possibly genetically determined, defect predisposing to the development of type 2 diabetes and evident prior to the onset of type 2 diabetes in high-risk individuals [222]. When ﬁrst-degree relatives of patients with type 2 diabetes were studied with the infusion of GIP during a hyperglycemic clamp experiment, a reduction of the insulinotropic GIP effect was found in ∼50% of the subjects [213]. However, from these experiments it was impossible to distinguish between a speciﬁc defect in the action of GIP and a general reduction of beta-cell function [223]. When the insulin secretory responses to a GIP bolus were tested at euglycemic conditions, the differences between ﬁrst-degree relatives and control subjects were no longer apparent [224]. Consistent with these studies, the secretion of GIP and GLP-1 as well as the overall contribution of the

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incretin effect to the postprandial rise in insulin secretion were found to be normal in ﬁrst-degree relatives of patients with type 2 diabetes [225,226]. Women with a history of gestational diabetes constitute another group at high risk for developing type 2 diabetes later in life, the empiric risk being on the order of 40%–70% [227]. However, in this particular cohort the insulinotropic effect of GIP was fully preserved both at euglycemia and under hyperclycemic clamp conditions and the secretion of both GIP and GLP-1 was not impaired [228]. Recently, Muscelli et al. examined the incretin effect in subjects with impaired glucose tolerance (IGT) [229]. In this study, the secretion of incretin hormones after an oral glucose load was similar in IGT subjects and controls, and there was no evidence for a speciﬁc defect in the beta-cell responsiveness towards GIP or GLP-1 [229]. There may, however, be slight alterations in GLP-1 secretion when a mixed meal rather than a pure glucose load is ingested, as suggested by Toft-Nielsen at al [203]. Thus, defects in the secretion of incretin hormones are unlikely to precede the development of type 2 diabetes [223]. Rather, an impairment in the betacell responsiveness to GIP develops alongside with a general decline in insulin secretion in subjects at high risk for type 2 diabetes. Likewise, disturbances in the postprandial secretion of GLP-1 may be a consequence rather than a cause of the metabolic abnormalities in type 2 diabetes.

Therapeutic Potential of Incretin Mimetics/Enhancers Owing to their potent glucose-lowering effects, both GIP and GLP-1 have been suggested as potential antidiabetic drugs [62,230].

Gastric Inhibitory Polypeptide Several groups have tested the effects of exogenous GIP administration in patients with type 2 diabetes. However, the insulinotropic effect of GIP is almost absent in these patients [208,210,212–214,231], and infusing GIP into hyperglycemic patients with type 2 diabetes has almost no effect on plasma glucose concentrations [209,232]. Moreover, unlike GLP-1, GIP has no effect on gastric emptying and at least in healthy volunteers stimulates glucagon secretion [60,199]. Taken together, these characteristics make GIP a rather unattractive drug candidate for the treatment of patients with diabetes [232]. The therapeutic administration of GIP is further limited by its short half-life in vivo. Thus, within ∼5–10 min, the intact peptide is cleaved by the enzyme dipeptidyl-peptidase 4 (DPP-4), yielding the inactive metabolite GIP(3–42) amide [233], which is eliminated via the kidneys with a half-life of ∼25 min [54]. A number of GIP analogues characterized by various amino acid

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exchanges or by the attachment of acyl residues have been generated in order to prevent the DPP-4 mediated degradation of the peptide [62,234–236]. It has been reported that some of these analogues exhibit a greater potency compared to native GIP in cell culture and animal experiments [62,232]. However, these chemical modiﬁcations do not prevent the GIP molecule from renal extraction and therefore cannot extend the half-life into a range typically required for once or twice daily administration [232]. Furthermore, it seems unlikely that the lack of efﬁcacy of GIP in patients with diabetes can be overcome by chemical modiﬁcations aiming to prolong the half life of the peptide.

Glucagon-Like Peptide-1 Unlike GIP, GLP-1 corrects hyperglycemia in patients with diabetes through several modes of action [215]. First, insulin secretion is enhanced by GLP-1 in a glucose-dependent manner [35,215]. This feature represents a major advantage over other insulinotropic drugs, such as sulfonylureas and benzoic acid derivatives, and provides a safeguard against the development of hypoglycemia [114]. It is also noteworthy that GLP-1 is still active in patients with type 2 diabetes after secondary failure to sulfonylurea treatment indicating that GLP-1 is capable of activating additional insulin secretory pathways that are not targeted by glucose or sulfonylureas [237]. One potential mechanism underlying the superior efﬁcacy of GLP-1 in patients with type 2 diabetes is the recruitment of insulin vesicles from the so called reserve pool into the readily releasable pool [238]. This mechanism allows for sustained provision of insulin secretory granules upon stimulation of the beta-cell. In addition, GLP-1 has been shown to increase insulin biosynthesis thereby preventing the chronic depletion of insulin granules [239]. Glucagon-like peptide-1 may also render beta cells previously insensitive to glucose stimulation glucoseresponsive [240]. Through these means, the percentage of functionally active beta cells may be increased by GLP-1 treatment. The second mechanism by which GLP-1 lowers glucose concentrations is suppression of glucagon secretion effectively counteracting the typical hyperglucagonemia of patients with diabetes [215,241]. The glucagonostatic effect of GLP-1 also allows for some reduction in glycemia in patients with type 1 diabetes [242,243]. While the GLP-1 effects on endocrine pancreatic secretion confer its glucose-lowering effects in the fasting state, the marked deceleration of gastric emptying is the primary mechanism for postprandial glucose control by GLP1 (vide supra) [79,127]. Thus, the typical postprandial rise in glucose and lipid concentrations is almost completely abolished with the administration of GLP-1 [79,127,244].

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While these properties make GLP-1 a near ideal drug candidate for the treatment of type 2 diabetes, its therapeutic application has been impeded by its unfavorable pharmacokinetic properties. Within ∼2 min after its intravenous or subcutaneous administration, intact GLP-1(7–36) amide is degraded to the metabolite GLP(9–36) amide [54,72,245], which has no direct effects on insulin and glucagon secretion or gastric emptying [168,170]. As a consequence, glucose control can only be achieved with native GLP-1 if the peptide is administered continuously, e.g., via a subcutaneous or intravenous infusion pump [143,246,247]. Two major strategies have been developed in order to circumvent these limitations. The ﬁrst one is to modify the structure of the GLP-1 molecule in order to prevent its degradation by DPP-4 and potentially delay its renal elimination [248]. Since DPP-4 degradation takes place at the N-terminal site of the peptide, most GLP-1 analogues display amino acid changes at the penultimate N-terminal position [248–250]. In addition, the biological half life of the peptide can be extended by the attachment of side chains inducing covalent or noncovalent binding to plasma albumin [123,250]. As a group, these GLP-1 based compounds-1 have been termed “incretin mimetics.” Due to the rapid progress in this ﬁeld, the major drug candidates currently available or under investigation will only brieﬂy be introduced.

Exenatide (Synthetic Exendin-4) The ﬁrst incretin mimetic to be introduced to the market was exenatide (Byetta®, Eli Lilly & Co.). This compound has been synthetically generated according to the amino acid sequence of the peptide exendin-4, which was discovered in the salivary gland of the Gila monster (a species of venomous lizard found in Arizona, USA) [251,252]. This peptide shares a 53% homology with the human GLP-1 molecule, which confers full activation of the GLP-1 receptor but resistance to DPP-4 in activation [251,252]. Therapeutic plasma levels of exendin-4 have been reached with twice daily subcutaneous administration [147–149,253]. A slow-release preparation of exendin-4 has been developed (AC2993 LAR, “long-acting release”), which might allow for glucose control with once weekly subcutaneous injection [254].

Liraglutide This GLP-1 analogue has been modiﬁed by an amino acid exchange as well as by the attachment of a fatty acid chain [250]. These changes induce DPP-4 resistance as well as noncovalent plasma albumin binding, thereby delaying renal ﬁltration and extending the in vivo half life to ∼11–15 h [255,256]. Other analogues of GLP-1 (e.g., albugon, CJC 1134) are currently undergoing clinical trials [257,258]. A detailed characterization of the pharmacokinetic proﬁles of these compounds is given elsewhere [123].

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As a rule, all biological actions of GLP-1 (i.e., stimulation of insulin secretion, suppression of glucagon secretion, deceleration of gastric emptying, inhibition of appetite, and stimulation of beta-cell regeneration) are mimicked by these incretin mimetics without major differences in their pharmacodynamics. The most important side effect of these drugs (as well as of native GLP-1) is the induction of nausea and (at higher plasma concentrations) vomiting [123]. These adverse reactions have been observed in up to 50% of the patients treated with incretin mimetics [147–149,256]. Typically, the nausea predominates during the ﬁrst few weeks of treatment, and the severity seems to be related to the respective dose administered [123]. It is not entirely clear whether the induction of nausea is linked to the deceleration of gastric emptying or whether this effect is directly mediated via GLP-1 receptors in the central nervous system (area postrema and circumventricular organ) projecting into the hypothalamus. However, usually the nausea disappears within the ﬁrst weeks after the initiation of treatment, and only a small percentage of patients have discontinued their therapy in clinical studies [123].

DPP-4 Inhibitors The second strategy to utilize the glucose-lowering potential of GIP and GLP1 for the treatment of diabetes is by blocking their degradation using speciﬁc DPP-4 inhibitors (also termed “incretin enhancers”), which can be taken orally [73,259]. By deﬁnition, the maximal efﬁcacy of such agents is limited by the extent of endogenous GLP-1 and GIP secretion. Given the modest rise in postprandial GLP-1 secretion [204], the increase in intact, biologically active GLP-1 levels (approximately twofold) achieved by DPP-4 inhibition appears relatively small [260]. This has led to some speculation as to whether the antidiabetogenic effects of these compounds might involve additional mechanisms other than the mere prolongation of GLP-1 activity [261]. Recent studies have demonstrated that not only DPP-4, but also neutral endopeptidase IV is capable of degrading the GLP-1 molecule [262]. Therefore, it appears worthwhile to combine inhibitors of both enzymes in order to achieve maximal effects. Since the DPP-4 inhibitors affect not only the levels of GLP-1 but also raise endogenous plasma levels of other bioactive peptides, especially of GIP [263,264], their effects differ from those of the incretin mimetics. Like the incretin mimetics, they lower glucose concentrations by stimulating insulin secretion and inhibiting glucagon secretion, with the latter mechanism predominating in some studies [260,265,266]. However, in contrast to the incretin mimetics, the DPP-4 inhibitors do not typically affect food intake or appetite [266]. Whether incretin enhancers also inhibit gastric emptying has

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not been studied to date, but appears unlikely. Since DPP-4 inhibitors rely on the endogenous secretion of GIP and GLP-1, they primarily affect postprandial glucose levels, whereas fasting glycemia is lowered to a lesser extent [260,265,266]. Despite some initial concerns that due to their lack of selectivity the DPP-4 inhibitors might inﬂuence other hormonal axes, these drugs are very well tolerated and so far no major adverse reactions have been reported with these drugs in clinical studies. A major advantage over the incretin mimetics is that nausea is not typically induced by the intake of DPP-4 inhibitors. The most developed compounds from this class of drugs are sitagliptin (Januvia®, Merck & Co.) and vildagliptin (Galvus®, Novartis Pharma). Reductions in HbA1c achieved during treatment with incretin mimetics were in the order of 0.8%–1.2%, and between 0.5% and 1.2% with DPP-4 inhibitors, depending on the doses and baseline HbA1c levels [147,256,260, 265,266]. A particular additional beneﬁt of the incretin mimetics is their weight-reducing effect, which appears to be consistent for the whole class and may progress over more than 100 weeks of treatment (with exenatide) so far [147–150].

Conclusion The study of physiological incretin hormones has shown that GIP and GLP-1 are important gut hormones that play a central role in maintaining normal glucose metabolism [267–269]. Impairment in their secretion and insulinotropic action may explain important facets of the pathophysiology of type 2 diabetes [15]. Furthermore, the unique characteristics of GLP-1, especially the well-preserved insulinotropic activity in type 2 diabetic patients and the glucose dependence of its action on insulin secretion, has stimulated the development of new therapies including the “incretin mimetics” and DPP-4 inhibitors.

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195. Larsen MO, Rolin B, Sturis J, Wilken M, Carr RD, Pørksen N, Gotfredsen CF. Measurements of insulin responses as predictive markers of pancreatic beta-cell mass in normal and beta-cell-reduced lean and obese Gottingen minipigs in vivo. Am J Physiol (Endocrinol Metab) 2006;290:E 670–677. Epub 2005 Nov 2008 196. Service GJ, Thompson GB, Service FJ, Andrews JC, Collazo-Clavell ML, Lloyd RV (2005) Hyperinsulinemic hypoglycemia with nesidioblastosis after gastric-bypass surgery. N Engl J Med 353:249–254 197. Meier JJ, Butler AE, Galasso R, Butler PC (2006) Hyperinsulinemic hypoglycemia after gastric bypass surgery is not accompanied by islet hyperplasia or increased beta-cell turnover. Diabetes Care 29:1554–1559 198. Tseng C-C, Kieffer TJ, Jarboe LA, Usdin TB, Wolfe MM (1996) Postprandial stimulation of insulin release by glucose-dependent insulinotropic peptide (GIP). Effect of a speciﬁc glucose-dependent insulinotropic polypeptide receptor antagonist in the rat. J Clin Invest 98:2440–2445 199. Meier JJ, Goetze O, Anstipp J, Hagemann D, Holst JJ, Schmidt WE, Gallwitz B, Nauck MA. Gastric inhibitory polypeptide does not inhibit gastric emptying in humans. Am J Physiol (Endocrinol Metab) 2004;286:E 621–625 200. Gallwitz B, Witt M, Fölsch UR, Creutzfeldt W, Schmidt WE (1993) Binding speciﬁcity and signal transduction of receptors for glucagon-like peptide-1(7–36) amide and gastric inhibitory polypeptide on RINm5F insulinoma cells. J Mol Endocrinol 10:259–268 201. Polonsky K, Rubenstein AH (1984) C-peptide as a measure of the secretion and hepatic extraction of insulin: pitfalls and limitations. Diabetes 33:486–493 202. Ross SA, Brown JC, Dupré J (1977) Hypersecretion of gastric inhibitory polypeptide following oral glucose in diabetes mellitus. Diabetes 26:525–529 203. Toft-Nielsen MB, Damholt MB, Madsbad S, Hilsted LM, Hughes TE, Michelsen BK, Holst JJ. Determinants of the impaired secretion of glucagon-like peptide-1 in type 2 diabetic patients. J Clin Endocrinol Metab 2001;86:3717–3723 204. Vilsbøll T, Krarup T, Deacon CF, Madsbad S, Holst JJ (2001) Reduced postprandial concentrations of intact biologically active glucagon-like peptide 1 in type 2 diabetic patients. Diabetes 50:609–613 205. Ranganath LR, Beety LM, Morgan LM, Wright JW, Howland R, Marks V (1996) Attenuated GLP-1 secretion in obesity: cause or consequence? Gut 38:916–919 206. Ebert R, Creutzfeldt W (1980) Hypo- and hypersecretion of GIP in maturity-onset diabetics. Diabetologia 19:271–272 207. Ørskov C, Jeppesen J, Madsbad S, Holst JJ (1991) Proglucagon products in plasma of noninsulin-dependent diabetics and nondiabetic controls in the fasting state and after oral glucose and intravenous arginine. J Clin Invest 87:415–423 208. Nauck MA, Heimesaat MM, Ørskov C, Holst JJ, Ebert R, Creutzfeldt W (1993) Preserved incretin activity of glucagon-like peptide 1 [7–36 amide] but not of synthetic human gastric inhibitory polypeptide in patients with type-2 diabetes mellitus. J Clin Invest 91:301–307 209. Amland PF, Jorde R, Aanderup S, Burhol PG, Giercksky K-E (1985) Effects of intravenously infused porcine GIP on serum insulin, plasma C-peptide, and pancreatic polypeptide in non-insulin-dependent diabetes in the fasting state. Scand J Gastroenterol 20:315–320 210. Jorde R, Burhol PG (1987) The insulinotropic effect of gastric inhibitory polypeptide in non-insulin dependent diabetes. Ital J Gastroenterol 19:76–78 211. Jones IR, Owens DR, Moody AJ, Luzio SD, Morris T, Hayes TM (1987) The effects of glucose-dependent insulinotropic polypeptide infused at physiological concentra-

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242. Creutzfeldt WO, Kleine N, Willms B, Ørskov C, Holst JJ, Nauck MA (1996) Glucagonostatic actions and reduction of fasting hyperglycemia by exogenous glucagon-like peptide I(7–36) amide in type I diabetic patients. Diabetes Care 19: 580–586 243. Dupré J, Behme MT, Hramiak M, McFarlane P, Williamson MP, Zabel P, McDonald TJ. Glucagon-like peptide 1 reduces postprandial glycemic excursions in IDDM. Diabetes 1995;44:626–630 244. Meier JJ, Gethmann A, Götze O, Gallwitz B, Holst JJ, Schmidt WE, Nauck MA. Glucagon-like peptide 1 abolishes the postprandial rise in triglyceride concentrations and lowers levels of non-esteriﬁed fatty acids in humans. Diabetologia 2006;49:1–7 245. Ritzel R, Ørskov C, Holst JJ, Nauck MA (1995) Pharmacokinetic, insulinotropic, and glucagonostatic properties of GLP-1 [7–36 amide] after subcutaneous injection in healthy volunteers. Dose-response-relationships. Diabetologia 38:720–725 246. Willms B, Idowu K, Holst JJ, Creutzﬂedt W, Nauck MA (1998) Overnight GLP-1 normalizes fasting but not daytime plasma glucose values in NIDDM patients. Exp Clin Edocrinol Diabetes 106:103–107 247. Rachman J, Gribble FM, Levy JC, Turner RC (1997) Near-normalization of diurnal glucose concentrations by continuous administration of glucagon-like peptide 1 (GLP-1) in subjects with NIDDM. Diabetologia 40:205–211 248. Gallwitz B, Ropeter T, Morys-Wortmann C, Mentlein R, Siegel EG, Schmidt WE (2000) Glucagon-like peptide-1-analogues resistant to degradation by dipeptidylpeptidase IV in vitro. Regul Pept 86:103–111 249. Gallwitz B, Schmidt WE, Conlon JM, Creutzfeldt W (1990) Glucagon-like peptide1(7–36)amide: characterization of the domain responsible for binding to its receptor on rat insulinoma RINm5F cells. J Mol Endocrinol 5:33–39 250. Knudsen LB, Nielsen PF, Huusfeldt PO, Johansen NL, Madsen K, Pedersen FZ, Thogersen H, Wilken M, Agersø H. Potent derivatives of glucagon-like peptide-1 with pharmacokinetic properties suitable for once daily administration. J Med Chem 2000;43:1664–1669 251. Eng J, Andrews PC, Kleinman WA, Singh L, Raufman JP (1990) Puriﬁcation and structure of exendin-3, a new pancreatic secretagogue isolated from Heloderma horridum venom. J Biol Chem 265:20259–20262 252. Eng J, Kleinman WA, Singh L, Singh G, Raufman JP (1992) Isolation and characterization of exendin-4, an exendin-3 analogue, from Heloderma suspectum venom. Further evidence for an exendin receptor on dispersed acini from guinea pig pancreas. J Biol Chem 267:7402–7405 253. Fineman MS, Bicsak TA, Shen LZ, Taylor K, Gaines E, Varns A, Kim D, Baron AD. Effect on glycemic control of exenatide (synthetic exendin-4) additive to existing metformin and/or sulfonylurea treatment in patients with type 2 diabetes. Diabetes Care 2003;26:2370–2377 254. Kim D, MacConnell L, Zhuang D, Scnabel C, Taylor K, Li W-I, Trautrmann M. Safety and efﬁcacy of a once-weekly, long-acting release formulation of exenatide over 15 weeks in patients with type 2 diabetes (abstract 487-P). Diabetes 2006;55 (Suppl. 1): A 116 255. Degn KB, Juhl CB, Sturis J, Jakobsen G, Brock B, Chandramouli V, Rungby J, Landau BR, Schmitz O. One week’s treatment with the long-acting glucagon-like peptide 1 derivative liraglutide (NN2211) markedly improves 24-h glycemia and alpha- and beta-cell function and reduces endogenous glucose release in patients with type 2 diabetes. Diabetes 2004;53:1187–1194

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256. Madsbad S, Schmitz O, Ranstam J, Jakobsen G, Matthews DR (2004) Improved glycemic control with no weight increase in patients with type 2 diabetes after oncedaily treatment with the long-acting glucagon-like peptide 1 analog liraglutide (NN2211): A 12-week, double-blind, randomized, controlled trial. Diabetes Care 27:1335–1342 257. Lawrence B, Wen SY, Jette L, Thibaudeau K, Castaigne J-P (2002) CJC-1131, the novel long-acting GLP-1 analogue, delays gastric emptying and demonstrates safety and tolerability in preclinical testing (abstract). Diabetes 51(Suppl 2):A84 258. Baggio LL, Huang Q, Brown TJ, Drucker DJ (2004) A recombinant human glucagonlike peptide (GLP)-1-albumin protein (albugon) mimics peptidergic activation of GLP-1 receptor-dependent pathways coupled with satiety, gastrointestinal motility, and glucose homeostasis. Diabetes 53:2492–2500 259. Deacon CF (2004) Therapeutic strategies based on glucagon-like peptide 1. Diabetes 53:2181–2189 260. Åhren B, Landin-Olsson M, Jansson PA, Svensson M, Holmes D, Schweizer A (2004) Inhibition of dipeptidyl peptidase-4 reduces glycemia, sustains insulin levels, and reduces glucagon levels in type 2 diabetes. J Clin Endocrinol Metab 89:2078–2084 261. Nauck MA, El-Ouaghlidi A (2005)The therapeutic actions of DPP-IV inhibition are not mediated by glucagon-like peptide-1 (for debate). Diabetologia 48:608–611 262. Plamboeck A, Holst JJ, Carr RD, Deacon CF (2005) Neutral endopeptidase 24.11 and dipeptidyl peptidase IV are both mediators of the degradation of glucagon-like peptide 1 in the anaesthetised pig. Diabetologia 48:1882–1890 263. Mentlein R, Gallwitz B, Schmidt WE (1993) Dipeptidyl-peptidase IV hydrolyses gastric inhibitory polypeptide, glucagon-like peptide-1(7–36)amide, peptide histidine methionine and is responsible for their degradation in human serum. Eur J Biochem 214:829–835 264. Mentlein R (1999) Dipeptidyl-peptidase IV (CD26)—role in the inactivation of regulatory peptides. Regul Pept 85:9–24 265. Åhren B, Gomis R, Standl E, Mills D, Schweizer A (2004) Twelve- and 52-week efﬁcacy of the dipeptidyl peptidase IV inhibitor LAF237 in metformin-treated patients with type 2 diabetes. Diabetes Care 27:2874–2880 266. Åhren B, Pacini G, Foley JE, Schweizer A (2005) Improved meal-related beta-cell function and insulin sensitivity by the dipeptidyl peptidase-IV inhibitor vildagliptin in metformin-treated patients with type 2 diabetes over 1 year. Diabetes Care 28:1936–1940 267. Scrocchi LA, Brown TJ, MaClusky N, Brubaker PL, Auerbach AB, Joyner AL, Drucker DJ. Glucose intolerance but normal satiety in mice with a null mutation in the glucagon-like peptide 1 receptor gene. Nat Med 1996;2:1254–1258 268. Hansotia T, Baggio LL, Delmeire D, Hinke SA, Yamada Y, Tsukiyama K, Seino Y, Holst JJ, Schuit F, Drucker DJ. Double incretin receptor knockout (DIRKO) mice reveal an essential role for the enteroinsular axis in transducing the glucoregulatory actions of DPP-IV inhibitors. Diabetes 2004;53:1326–1335 269. Miyawaki K, Yamada Y, Yano H, Niwa H, Ban N, Ihara Y, Kubota A, Fujimoto S, Kajikawa M, Kuroe A, Tsuda K, Hashimoto H, Yamashita T, Jomori T, Tashiro F, Miyazaki J-i, Seino Y. Glucose intolerance caused by a defect in the entero-insular axis: A study in gastric inhibitory polypeptide receptor knockout mice. Proc Natl Acad Sci USA 1999;96:14843–14847 270. Buffa B, Polak JM, Pearse AGE, Solcia E, Grimelius L, Capella C (1975) Identiﬁcation of the intestinal cell storing gastric inhibitory polypeptide. Histochemistry 43:249– 255

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271. Buteau J, Foisy S, Rhodes CJ, Carpenter L, Biden TJ, Prentki M (2001) Protein kinase Czeta activation mediates glucagon-like peptide-1-induced pancreatic beta-cell proliferation. Diabetes 50:2237–2243 272. Gromada J, Bokvist K, Ding WG, Holst JJ, Nielsen JH, Rorsman P (1998) Glucagonlike peptide 1 (7–36) amide stimulates exocytosis in human pancreatic beta cells by both proximal and distal regulatory steps in stimulus-secretion coupling. Diabetes 47(1):57–65

V. Pancreatic Beta Cell and Disease

18. Pancreatic Islet Pathology in Type 2 Diabetes Anne Clark

Summary. Inadequate insulin secretion is a major factor in type 2 diabetes. Islet dysfunction is evident at the onset of the disease but the causal factors are largely unknown; decreased beta-cell mass, as shown in rodent models of diabetes, has been proposed. Quantitative morphometry of post-mortem human pancreas has demonstrated 0–50% less beta-cell population in diabetic compared to nondiabetic subjects. The evidence for continuous turnover of beta cells in adult human pancreas by apoptosis and regeneration (as in rodents) has yet to be proven; low incidence of cell division in ducts or islets has been reported and no evidence for abnormal cell turnover contributing to decreased beta-cell mass in human diabetes has been seen. The functional capacity for increased insulin secretion is high in diabetic and nondiabetic subjects. It is likely that a deﬁcit in function is a major contributor for type 2 diabetes in man. Islet amyloid deposition is heterogeneous in human diabetes and is unlikely to contribute to decreased beta-cell mass at onset of hyperglycemia in man as shown in rodent models, monkeys, and cats. The effects of elevated lipids on beta-cell function is mediated by modulation of metabolism but the cytotoxic effects of lipids on beta cells—lipotoxicity—is mediated largely by aberrant effects of saturated fatty acids on cells in vitro. Increased availability of human isolated islets will enable accurate determination of functional and morphological changes which contribute to impaired insulin secretion in type 2 diabetes.

Introduction The role of pancreatic islets in the onset and progression of type 2 diabetes is of major signiﬁcance. However, pathological changes in human islets which contribute to the aberrant pattern and eventual loss of insulin secretion in Diabetes Research Laboratories, Oxford Centre for Diabetes, Endocrinology and Metabolism, Churchill Hospital, Oxford OX3 7LJ, UK

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type 2 diabetes have been largely inferred from pancreatic specimens taken at post mortem. Increased availability of islets from human donors (including some from diabetic subjects) has allowed more detailed functional and morphological examination of pathophysiology. It is now apparent that pathological changes associated with hyperglycemia are not identical in human and rodent islets. Thus, direct extrapolation from rodents to man of diseaserelated pathophysiology must be made with caution. Currently, there is a debate concerning the relative roles in the etiology of type 2 diabetes of decreased beta-cell mass compared with decreased beta-cell function and the interdependence of these two factors [1–4]. The potential for regeneration and replacement of islet cells in adults is of signiﬁcance for both type 1 and type 2 diabetes. Islet amyloidosis and the associated loss of endocrine cells is found in human type 2 diabetes but not in rodents; the role of amyloid in the onset and progression of type 2 diabetes is unclear. Furthermore, identiﬁcation of susceptibility genes for type 2 diabetes, particularly those of MODY [5,6], has focused attention on factors associated with function and embryological development of islet cells [7–10]; these factors could also be important in susceptibility for type 2 diabetes. In addition, metabolic factors, such as hyperlipidemia, are thought to have a role in the lipotoxicity associated with type 2 diabetes and could affect both viability and function of islet cells [11].

The Role of Beta-Cell Mass in the Etiology of Type 2 Diabetes Islets in patients with type 2 diabetes contain a large population of beta cells (40%–80% of islet mass) and appear morphologically similar to islets from similarly-aged nondiabetic subjects; the only microscopically abnormal feature in humans is the development of amyloid in some islets [12–14]. Whereas the acute reduction in insulin secretion in type 1 diabetes is related to a severe decline or total loss of the population of beta cells [15], this is not the situation in most patients with type 2 diabetes. It is therefore unclear if the insulin secretion which is inadequate to maintain normoglycemia at the onset of type 2 diabetes in man results from a deﬁcit of function (e.g., in stimulus-secretion coupling mechanisms for insulin and glucagon secretion), or a decreased mass of otherwise normally functioning beta cells or a combination of both [16]. The proportions of beta cells in patients with established type 2 diabetes have been reported to be unchanged or reduced; published data from analysis of post-mortem tissue indicate either no change or 6%–54% less beta cells in comparison with nondiabetic subjects [17–25] (Fig. 1). The variability of

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Differences in % β-cell/pancreas in Type 2 diabetes compared to controls

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Fig. 1. Comparison of published data on differences of beta-cell proportions in the pancreas of diabetic and nondiabetic subjects. The beta-cell proportion was calculated in relation to the exocrine pancreas. Data was derived from published morphometric observations on post-mortem pancreatic material from diabetic and nondiabetic subjects [18,20–25,110] and expressed as a percentage mean difference between the diabetic and nondiabetic groups. For comparative purposes, data reported separately for obese and lean subjects by Butler in 2003 [25] was combined. The difference in beta-cell proportion varied between no difference between the groups in three studies to a maximum of 54% less beta-cell proportion in diabetes

these published data reﬂect not only the heterogeneous nature of the disease but also problems inherent in analyses of human post-mortem tissue. Specimens have been taken predominantly from patients who have had many years of diabetes and it is therefore difﬁcult to predict which changes have arisen as a result of the diabetes rather than were causal for the disease. Often, only small tissue samples are available. This means that an inadequate representation of the approximately one million islets per pancreas can be examined. In addition, quantitative morphometry of histological sections is made such that islet features are measured in relation to the exocrine tissue or to pancreatic weight or volume; exocrine tissue can also be affected by disease, e.g., exocrine ﬁbrosis due to chronic pancreatitis [26] or fat inﬁltration in obesity [21,27,28]. Recent studies on post-mortem pancreas from glucose intolerant subjects are more likely to reﬂect the beta-cell population in patients at risk for onset of diabetes and at onset of hyperglycemia [25,29]. A smaller population of beta cells (40% less than present in nondiabetic controls) was found in subjects with blood glucose levels in the range >6.1 < 7.2 mM; however, the wide distribution of these data show that there is considerable overlap of beta-cell

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/exocrine tissue proportion between the impaired fasting glucose (IFG) (0.5– 3.7 range; median 0.8%) and normoglycemic groups (0.65–11.0; median 2%) [29]. A more extensive analysis of a larger number of islets/pancreas than examined by the Butler group [23] indicated that the islet density in the pancreas was not different between diagnosed diabetic and control subjects and there was 33% fewer beta cells/pancreas in patients with established diabetes compared to controls [23]. These authors concluded that such a difference in the beta-cell population would be unlikely to be a major factor in the onset of hyperglycemia in type 2 diabetes in Japanese subjects; insulin-treated patients had a lower beta-cell volume compared to those treated with oral agents, suggesting a progressive deterioration with disease [23,28,30]. Thus, from morphological studies, it is unclear if onset of hyperglycemia results from insufﬁcient beta-cell mass or a deﬁcit in islet function or a combination of both. Many factors determine the size of the beta-cell population in normal subjects. Islet cell development is governed by transcription factors [8,31] and could be modiﬁed by genetically determined alterations in these factors some of which regulate not only islet development but also insulin and glucagon gene expression, e.g., PDX-1, NeuroD1/Beta2, PAX6 [32]; mutations in these genes are associated largely with MODY [6]. In addition environmental factors associated with increased demand for insulin secretion such as increased nutritional stimulation, pregnancy, and obesity modulate the adult beta-cell population in rodents [33,34]. Human islets exhibit plasticity of function but little is known about ﬂuctuations in their mass [35]. Thus the “normal” population of beta cells in humans could be very heterogeneous. It is difﬁcult to see how a reduction of less than 50% of otherwise healthy beta cells could induce hyperglycemia, since a recent in vivo organ donation of islets from 50% of a healthy human pancreas did not induce hyperglycemia in the donor and also established normoglycemia in the previously diabetic recipient [36]. In dogs, 60% pancreatectomy did not induce hyperglycemia [37], although a 50% pancreatectomy induced IFG and a modiﬁcation of the pattern of insulin secretion [38]. However, rodents are the species in which most studies have been done and, in otherwise healthy rats and mice, a 95% pancreatectomy is required to induce diabetes. It therefore seems unlikely that a 40–50% reduction in healthy functional beta cells could be the sole cause of IFG in man. Under normal conditions, the beta-cell population exceeds that required to maintain normoglycemia in most species. In man, it is estimated that there are one million islets in a normal adult pancreas and that less than 0.5% of the pancreatic reserve of insulin granules is required for ﬁrst phase insulin secretion following a glucose challenge [39]. In increased insulin resistance, when fasting and postprandial beta-cell secretion can be more than doubled and during acute exercise or in pregnancy, beta-cell secretion is automatically

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increased [35]. Beta-cell hyperplasia and increased islet size has been reported in pregnant rodents [40] and some similar observations made in post-mortem tissue in pregnant humans [41]. However, less than 12% of obese individuals with insulin resistance develop type 2 diabetes [42], suggesting efﬁcient functional compensation of insulin secretion in most individuals. Moreover, insulin secretion can be more than doubled in a very short time in type 2 diabetic subjects by infusions of GLP-1 [43] and ﬁrst-phase insulin secretion can be partially restored by sulfonylureas [44]. These observations taken together indicate that there is a large reserve capacity for insulin production in human pancreas and that a moderate reduction in beta-cell mass would not be important unless function in terms of nutrient-stimulated insulin secretion was also compromised [16].

Regeneration and Turnover of the Beta-Cell Population It has been suggested from studies in rodents that diabetes results from an imbalance of a normal cycle of islet cell turnover [34,45]. It has been thought for many years that, similar to the life cycle of most neurons, the islet cell population develops in utero and is fully established probably by the end of the second decade and that this cell population remains for the lifetime of the individual. However, a proposal has been made from studies in rodents that there is a program of continuous islet cell turnover—death by apoptosis and regeneration by neogenesis from ductal precursor cells or division of existing beta cells [46]. Thus, a deﬁcit of islet beta cells could result from either an increase in apoptosis or a decreased neogenesis of beta cells [47,48]. At present, there is less evidence for extensive regeneration of beta cells in human pancreas compared to that found in rodents; quantitative observations on neogenesis from ductal stem cells, beta-cell division, or death by apoptosis in normal human islets have been made largely on post-mortem pancreas and the degrees of cell division or cell death are generally very low [25,49]. In adult human pancreas, isolated beta cells or small groups of endocrine cells are found adjacent to cytokeratin 19 positive pancreatic ductal cells (Fig. 2) and these extra-islet cells can form up to 15% of the total population of beta cells [4,49]. The number of these putative neogenic cells is reported to be unchanged [25] or increased [4] in type 2 diabetes. It has been assumed from their localization that these cells represent recent neogenesis from ductal cell precursors [47] as occurs in islet cell embryogenesis. However, the classiﬁcation of these cells as neogenic is uncertain; the age of the cells (weeks/ months/years) has not been determined nor has their potential to form new islets or their functional capacity for appropriate insulin secretion. Ductal

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Fig. 2. Localization of beta cells in islets, clusters, and adjacent to exocrine ducts in nondiabetic human pancreas. Pancreatic specimens from a nondiabetic adult subject showing insulin-positive cells (red) localized by immunocytochemistry in islets and in small clusters (Cl) as well as single cells adjacent to ductal epithelia (D) (cytokeratin 19, CK19+ positive, brown) (arrows indicate ductal beta cells). Scale bar: 50 µm

endocrine cells are increased in pancreatic pathology such as adenocarcinomas and in pancreatitis [50], suggesting that there is potential for neogenesis of islet cells in adults but the stimulus for their proliferation is unknown. If these cells are to be considered to be neogenic and an important source of “new” islets, their functional capacity for hormone secretion should be determined. This is especially important in view of the current proposal that some diabetes treatment regimes e.g., GLP-1 are mediated by increased beta-cell neogenesis, as demonstrated in rodents [51,52]. An alternative system for beta-cell expansion has been identiﬁed in established rodent islets by division of differentiated beta cells [46]; this results in increased islet size but no change in islet number; islet size in rodents increases in response to increased insulin requirement in obesity, e.g., ob/ob mice [53], hyperglycemia [54], and with increase in body weight [55]. In man, the fre-

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quency of beta-cell replication (Ki67 positive) is very low in adults and there is no evidence for increased replication in obesity [25]. Increased beta-cell mass has been described in European obesity [28] but no relationship was found between islet size and body mass in a recent study of Asian obese and type 2 diabetic subjects [24], and no change in islet size was seen between obese and lean nondiabetic or diabetic subjects [25]. The fact that turnover of islet cells (replication and apoptosis) is very low in aged mice (>9 months old) [56] suggests that young rodents may not be the best example for comparisons with adult man.

Islet Amyloid and Type 2 Diabetes Islet amyloidosis is a characteristic feature of patients with type 2 diabetes (Fig. 3) and was one of the ﬁrst characteristics to separate juvenile-onset and maturity-onset disease [57]. Islet amyloidosis is associated with loss of beta cells but the degree of amyloid ﬁbril-induced beta-cell loss is very heterogeneous in man. At post-mortem, more than 90% of type 2 diabetic subjects have amyloid deposits in at least one islet and the deposits are less frequently found (up to 12%) in nondiabetic subjects of similar age [13,30,58]. Islet amyloidosis is a feature of diabetic populations worldwide including the Japanese [18,23], North American Pima Indians [59], and Asians [24,60,61]. The distribution of islet amyloid is heterogeneous both within a pancreas and in different subjects [62]. The prevalence (number of islets affected) can vary between <1% to >80% in patients with established diabetes. The degree of amyloidosis (percentage of islet containing amyloid) is also variable but there is a positive association between the degree and prevalence of islet amyloid [12,63,64] (Fig. 4). Amyloid deposition is progressive and develops initially adjacent to islet capillaries (Fig. 3). As the deposits become more extensive, the ﬁbrils replace endocrine cells and inﬁltrate to ﬁll larger islet spaces with more islets becoming affected [65] (Fig. 4). There is no relationship between extent of islet amyloidosis and duration of diabetes, but patients with more severe islet dysfunction requiring insulin replacement therapy have, at postmortem, more extensive amyloidosis [30]. This implies that extensive amyloidosis is associated with decline in insulin secretion in the course of the diabetic syndrome. However, there is no evidence that amyloid is a major factor for the onset of hyperglycemia in man; some patients with diabetes of many years’ duration have less than 1% of their islets affected suggesting that amyloid deposition is a complication rather than an etiological factor for the pathogenesis of the disease [66,67]. Islet amyloid ﬁbrils are formed from islet amyloid polypeptide (IAPP), also known as “amylin” [68–70], which is a normally occurring peptide of the beta

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B

A

Amyloid

C BM

β −cell

Monkey D

β −cell

hIAPP TM

amy BM cap

amy cap

Fig. 3. Islet amyloid deposits in pancreatic islets of A,B diabetic human subjects, C a diabetic monkey, and D a nondiabetic transgenic mouse expressing the gene for human islet amyloid polypeptide (hIAPP TM). Islet amyloid ﬁbrils (labeled green with ﬂuorescent thioﬂavin S in (A and B) are deposited initially adjacent to islet capillaries (A) but progress to largely ﬁll the islet space (B) associated with destruction of islet endocrine cells. The degree of amyloidosis is very heterogeneous both within the pancreas and between diabetic subjects. Perivascular amyloid (amy) visualized by electron microscopy (C and D) appears as an amorphous material, immunogold labeled for IAPP situated between the basement membrane (BM) of the capillary (cap) and the beta cells. This could functionally impair signaling between the circulation and islets cells and hormone release into the circulation. Scale bars: A,B 20 µm; C 5.0 µm; D 1.0 µm

cell, co-stored with insulin in secretory granules and co-released in response to beta-cell secretagogues [71,72]. Genetic alterations in the coding and promoter regions of the IAPP gene have been linked to diabetes in very small numbers of diabetic subjects in Japan [73], China [61], Spain [74], and Polynesia [75]. However, there is no evidence for genetically determined abnormalities of the IAPP gene in the majority of patients with type 2 diabetes [76–78]. It is therefore likely that abnormal protein folding of this highly hydrophobic peptide results from some environmental change in the region of the islet. The causal factors for refolding of IAPP into a beta-sheet conformation and molecular oligomerization to form amyloid ﬁbrils are largely unknown; over-

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100

% severity

80 60

R2 = 0.6388

40 20 0 0

20

40

60

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100

% prevalence

Fig. 4. The degree of islet amyloidosis in 39 diabetic patients determined by quantitation of thioﬂavin S-labeled pancreatic islets in post-mortem tissue sections. The severity (degree of islet area occupied by amyloid) and the prevalence (number of islets affected) are positively associated (R2 = 0.64), indicating that as the severity of the amyloid progresses more islets are affected. More than 50% of the subjects (n = 20) with established diabetes of many years’ duration have less than 20% of islets affected with less than 10% of islet mass occupied with amyloid (arrows)

production and decreased clearance of the peptide from extracellular islet spaces have been proposed. Currently, the best hypothesis for diabetes-related islet amyloidosis is secretion of incompletely processed precursor peptide, proIAPP. ProIAPP fragments have been identiﬁed in islet amyloid deposits [79] and proIAPP binds to heparan sulfate, which is a basement membrane proteoglycans and has itself been shown to accelerate IAPP ﬁbril formation [80]. Inhibition of proteolysis of proIAPP by the hormone convertase enzyme, PC2, accelerates ﬁbril formation in cellular and mouse models [81,82]. Increased production of incompletely processed proinsulin is a characteristic feature of type 2 diabetes [83]. Thus in type 2 diabetes, increased proinsulin secretion could be accompanied by increased proIAPP and initiation of ﬁbrillogenesis. Whilst islet amyloid is unlikely to be a causal factor for onset of hyperglycemia in type 2 diabetes in man, it appears to be a major factor for induction of diabetes in animal models of amyloid-associated diabetes. Islet amyloidosis progressively replaces islet beta cells in diabetic monkeys [84] and transgenic mice expressing the gene for human IAPP (hIAPP TM) [85], and the degree of amyloidosis is related to onset of glucose intolerance in hIAPP TM and in spontaneous diabetes in cats [65,86]. Therapies which improve glucose intolerance and reduce IAPP secretion can reduce the deleterious effects of islet amyloid in these models [87,88].

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Differences between the animal models and man indicate that factors other than amyloid-induced beta-cell death must contribute to onset of hyperglycemia in man. However, replacement of endocrine cells with amyloid is undoubtedly associated with severe decline of islet function and requirement for insulin therapy, which occurs later in the diabetic syndrome. The exact mechanisms by which amyloid ﬁbrils are cytotoxic in vivo are unclear; synthetic human IAPP ﬁbrils in vitro are toxic to cells by induction of apoptosis [82], but it may not be the mature ﬁbrils that induce cell death but an early intermediate of ﬁbrillogenesis such as small oligomers [89].

Lipotoxicity The concept of lipotoxicity has developed as a result of the interest in the effects of lipids and, in particular, nonesteriﬁed fatty acid (NEFAs or fatty acids) on islet cell secretion. Pioneering work by Denis McGarry showed that effective islet cell secretion required NEFAs to be present as a substrate for metabolism [90–93]. Acute administration of NEFAs (over hours rather than days) increases glucose stimulated insulin secretion (GSIS) both in vivo in man [94] and in vitro [95]. These observations have been conﬁrmed and extended by in vitro observations demonstrating that NEFAs increase insulin secretion by promoting closure of K+ATP channels [96], increased conversion to fatty acid CoA, beta-oxidation, and increased ATP production and/or fatty acid/ triglyceride (TG) cycling [97,98]. However, long-term exposure of islets or clonal cells to NEFAs results in decreased insulin production and induction of so-called “lipotoxicity” [92,98,99]. The mechanisms for this deleterious effect of NEFAs remain unclear. Chronic exposure of islet cells to lipids decreases insulin gene expression and has been postulated to induce apoptosis and cell death—“lipotoxicity” [100,101]. Rodent models for these studies have been the Zucker diabetic fatty rat (ZDF) and ZF rat, which are models of hyperphagia as a result of a leptin receptor mutation. In some ZDF animals, there is development of obesity and hyperlipidemia, and failure to compensate by adequate insulin secretion is associated with decreased islet cell population and onset of diabetes [11,102]. It has been assumed that this was a good model for obesity-related, type 2 diabetes in man and the role of lipotoxicity. However, in man fewer than 12% of obese hyperlipidemic subjects become diabetic [42,103] and there is no evidence for a reduction in the beta-cell mass in obesity. The degree of lipidinduced apoptosis and reduction in glucose-stimulated insulin secretion from beta cells in vitro was found to be greater with saturated compared to unsaturated NEFAs [95,104]. Lipid-induced cytotoxicity was shown to result from increases in endoplasmic stress (ER stress), ceramide production and

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induction of genes that can result in apoptosis [105,106]. However, in vitro the saturated NEFA, palmitic acid (which has been the NEFA of choice) is converted to the insoluble triglyceride, tripalmitin, in cultured cells and this rigid accumulation in the ER results in cell death [107]. More importantly, in vivo there is always a mixture of saturated and unsaturated NEFAs in both normal and elevated lipid conditions so that tripalmitin would never be formed. Thus lipotoxic effects inducing cell death in vitro are largely a result of artifactual manipulation of NEFAs and formation of an abnormal intracellular TG. It is indisputable that chronic exposure of beta cells to elevated concentrations of NEFAs result in metabolic disturbances and reduction of glucosestimulated insulin secretion, even though this does not lead inevitably to diabetes. This chronic exposure to lipids could result from local, but not necessarily intracellular lipid stores. Unlike clonal cells in vitro, islets from obese humans or mice do not contain large droplets of lipid stored as TG. Pancreatic fat in obese man is elevated in the form of extensive swollen adipocytes in the exocrine tissue (Fig. 5). Lipid fractions will be exchanged between the adipocytes and the circulation via the extracellular space.

A

Exo

B

Islet Islet Exo

Ad Ad

Islet

Fig. 5A,B. Localization of lipid stores within the pancreas. A Post-mortem pancreas from an obese diabetic subject showing swollen adipocytes (Ad) within the exocrine tissue (Exo) and close to an islet. B Localization of a pancreatic islets (immunolabeled for insulin) adjacent to swollen adipocytes (Ad) in the exocrine tissue (Exo) of an obese patient. Products of lipid metabolism would be elevated in the extracellular space adjacent to islets. Scale bars: 50 µm

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Adjacent islet cells would then be exposed to high levels of extracellular NEFAs which could, in susceptible individuals, result in abnormal regulation of plasma glucose. Human islet isolation for transplantation has provided opportunities to examine both structural and functional aspects of human isolated islets from nondiabetic and diabetic subjects. This will enable more direct comparisons of details of stimulus-secretion coupling, gene proﬁling, metabolism, and oxidative damage in islets from diabetic and nondiabetic subjects [108,109]. Preliminary reports of increased apoptosis and decreased mitochondrial function in human isolated islets from diabetic subjects have been made [109]. As more of these islets become available for laboratory investigation, it will be possible to identify exact determinants of impaired islet function which contribute to islet cell dysfunction in type 2 diabetes in man. Acknowledments. I am grateful to students and members of my research team for their contributions and to Professor John Morris in the Department of Human Anatomy and Genetics for studies with the electron microscope. Funding for some of this work was provided by the Wellcome Trust, UK.

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84. de Koning EJ, Bodkin NL, Hansen BC and Clark A (1993) Diabetes mellitus in Macaca mulatta monkeys is characterised by islet amyloidosis and reduction in beta-cell population. Diabetologia 36:378–384 85. Janson J, Soeller WC, Roche PC, Nelson RT, Torchia AJ, Kreutter DK, Butler PC (1996) Spontaneous diabetes mellitus in transgenic mice expressing human islet amyloid polypeptide. Proc Natl Acad Sci U S A 93:7283–7288 86. O’Brien T, Hayden D, Johnson K and Fletcher T (1986) Immuno-histochemical morphometry of pancreatic endocrine cells in diabetic normoglycaemic glucoseintolerant and normal cats. J. Comp. Pathol. 96:357–369 87. Hull RL, Shen ZP, Watts MR, Kodama K, Carr DB, Utschneider KH, Zraika S, Wang F, Kahn SE (2005) Long-term treatment with rosiglitazone and metformin reduces the extent of, but does not prevent, islet amyloid deposition in mice expressing the gene for human islet amyloid polypeptide. Diabetes 54:2235–2244 88. Hoenig M, Hall G, Ferguson F, Jordan K, Henderson M, Johnson K, O’Brien T (2000) A feline model of experimentally induced amyloidosis. American Journal of Pathology. 157:2143–2150 89. Meier JJ, Kayed R, Lin CY, Gurlo T, Haataja L, Jayasinghe, S, Langen P, Glabe CC, Butler PC (2006) Inhibition of human IAPP ﬁbril formation does not prevent betacell death: evidence for distinct actions of oligomers and ﬁbrils of human IAPP. Am J Physiol Endocrinol Metab 291:E1317–1324 90. Dobbins RL, Chester MW, Stevenson BE, Daniels MB, Stein DT, McGarry JDl (1998) A fatty acid- dependent step is critically important for both glucose- and nonglucose-stimulated insulin secretion. J Clin Invest 101:2370–2376 91. Stein DT, Esser V, Stevenson BE, Lane KE, Whiteside JM, Daniels MB Chen S, McGarry JD (1996) Essentiality of circulating fatty acids for glucose-stimulated insulin secretion in the fasted rat. J Clin Invest 97:2728–2735 92. Prentki M and Corkey BE (1996) Are the beta-cell signaling molecules malonyl-CoA and cystolic long-chain acyl-CoA implicated in multiple tissue defects of obesity and NIDDM? Diabetes 45:273–283 93. Corkey BE, Deeney JT, Yaney GC, Tornheim K and Prentki M (2000) The role of long-chain fatty acyl-CoA esters in beta-cell signal transduction. J Nutr 130: 299S–304S 94. Beysen C, Karpe F, Fielding BA, Clark A, Levy JC, Frayn KN (2002) Interaction between speciﬁc fatty acids, GLP-1 and insulin secretion in humans. Diabetologia 45:1533–1541 95. Gravena C, Mathias PC and Ashcroft SJ (2002) Acute effects of fatty acids on insulin secretion from rat and human islets of Langerhans. J Endocrinol 173:73–80 96. Branstrom R, Leibiger IB, Leibiger B, Corkey BE, Berggren PO, Larsson O (1998) Long chain coenzyme A esters activate the pore-forming subunit (Kir6. 2) of the ATPregulated potassium channel. J Biol Chem 273:31395–31400 97. Larsson O, Deeney JT, Branstrom R, Berggren PO and Corkey BE (1996) Activation of the ATP-sensitive K+ channel by long chain acyl-CoA. A role in modulation of pancreatic beta-cell glucose sensitivity. J Biol Chem 271:10623–10626 98. Roduit R, Nolan C, Alarcon C, Moore P, Barbeau A, Delghingaro-Augusto V, Przybykowski E, Morin J, Masse F, Massie B, Rudenramn N, Phodes C, Pouitout V, Prentki M (2004) A role for the malonyl-CoA/long-chain acyl-CoA pathway of lipid signaling in the regulation of insulin secretion in response to both fuel and nonfuel stimuli. Diabetes 53:1007–1019 99. Unger RH and Orci L (2002) Lipoapoptosis: its mechanism and its diseases. Biochim Biophys Acta 1585:202–212

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100. Cnop M, Hannaert JC, Hoorens A, Eizirik DL and Pipeleers DG (2001) Inverse relationship between cytotoxicity of free fatty acids in pancreatic islet cells and cellular triglyceride accumulation. Diabetes 50:1771–1777 101. Briaud I, Rouault C, Reach G and Poitout V (1999) Long-term exposure of isolated rat islets of Langerhans to supraphysiologic glucose concentrations decreases insulin mRNA levels. Metabolism 48:319–323 102. Unger RH and Zhou YT (2001) Lipotoxicity of beta-cells in obesity and in other causes of fatty acid spillover. Diabetes 50 Suppl 1:S118–121 103. Ford ES, Giles WH and Dietz WH (2002) Prevalence of the metabolic syndrome among US adults: ﬁndings from the third National Health and Nutrition Examination Survey. Jama 287:356–359 104. Maedler K, Oberholzer J, Bucher P, Spinas GA and Donath MY (2003) Monounsaturated fatty acids prevent the deleterious effects of palmitate and high glucose on human pancreatic beta-cell turnover and function. Diabetes 52:726–733 105. Robertson RP, Harmon J, Tran PO, Tanaka Y and Takahashi H (2003) Glucose toxicity in beta-cells: type 2 diabetes, good radicals gone bad, and the glutathione connection. Diabetes 52:581–587 106. Kharroubi I, Ladriere L, Cardozo AK, Doguso Z, Cnop M, Eisirik DL (2004) Free fatty acids and cytokines induce pancreatic {beta}-cell apoptosis by different mechanisms: role of NF-{kappa}B and endoplasmic reticulum stress. Endocrinology 107. Mofﬁtt JH, Fielding BA, Evershed R, Berstan R, Currie JM, Clark A (2005) Adverse physicochemical properties of tripalmitin in beta cells lead to morphological changes and lipotoxicity in vitro. Diabetologia 48:1819–1829 108. Deng S, Vatamaniuk M, Huang X, Doliba N, Lian MM, Frank A, Velidideoglu E, Desai NM, Koberlein B, Wolf B, Braker CF, Naji A, Matchinsky FM, Markmann JF (2004) Structural and functional abnormalities in the islets isolated from type 2 diabetic subjects. Diabetes 53:624–632 109. Marchetti P, Del Guerra S, Marselli L, Lupi R, Masini M, Pollera M, Bughani H, Boggi U, Vistoli F, Mosca F, Del Prato S (2004) Pancreatic islets from type 2 diabetic patients have functional defects and increased apoptosis that are ameliorated by metformin. J Clin Endocrinol Metab 89:5535–5541 110. Stephan Y, Orci L, Malaisse-Lagae F, Perrelet A, Patel Y, Unger RH (1982) Quantitation of endocrine cell content in the pancreas of non-diabetic and diabetic humans. Diabetes 31:694–700

19. Genetic Disorders of the Pancreatic Beta Cell and Diabetes (Permanent Neonatal Diabetes and Maturity-Onset Diabetes of the Young) Emma L. Edghill and Andrew T. Hattersley

Summary. Mutations in critical beta-cell genes can result in monogenic diabetes. This clinically heterogeneous group of disorders usually presents soon after birth as neonatal diabetes, or during childhood or early adulthood as maturity-onset diabetes of the young (MODY). Most defects arise in genes involved in pancreatic beta-cell development or the maintenance of beta-cell function. Studying the phenotype of patients with mutations and the mechanisms by which these mutations result in diabetes gives new insights into normal and pathological functioning of the beta cell. The most common genetic etiology in patients with MODY are mutations in the genes that encode the transcription factors hepatocyte nuclear factor (HNF)-1 alpha (TCF1), HNF-4 alpha (HNF4A) and HNF-1 beta (TCF2), and the glycolytic enzyme glycokinase (GCK). Mutations in each of these genes result in different clinical phenotypes and cause beta-cell dysfunction through different mechanisms. The commonest causes of neonatal diabetes are defects in betacell function, arising from mutations in genes encoding the subunits which form the KATP channel, Kir6.2 (KCNJ11) and SUR1 (ABCC8). Deﬁning the genetic subtypes of monogenic diabetes not only helps understanding of the beta cell, it also has considerable implications for patient care. A genetic diagnosis provides accurate information regarding inheritance, prognosis, can explain clinical features and may guide patient treatment. The best example of pharmacogenetics is that patients with KCNJ11 mutations, despite being insulin dependent, can have excellent glycemic control on Institute of Biomedical and Clinical Science, Peninsula Medical School, Barrack Road, Exeter, Devon EX2 5AX, UK

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high-dose sulfonylureas. Deﬁning the genetic etiology of monogenic diabetes has therefore contributed both to science and patient care.

Introduction to Monogenic Diabetes Monogenic diabetes is a group of single gene disorders where a mutation or mutations in a single gene result in diabetes. It is estimated that 1%–2% of all patients with diabetes have monogenic diabetes although they are frequently not diagnosed. The vast majority of monogenic diabetes results from beta-cell dysfunction. The usual presentation of mutations causing beta-cell defects are permanent neonatal diabetes (PNDM), transient neonatal diabetes (TNDM), maturity-onset diabetes of the young (MODY), or genetic syndromes. This chapter will focus on PNDM and MODY, describing their known genetic etiologies, clinical characteristics and proposed pathophysiology.

Clinical Classifications of Neonatal Diabetes Neonatal diabetes is a rare form of diabetes diagnosed in early infancy with a prevalence of approximately 1 in 1–200 000 births. Diabetes diagnosed throughout the ﬁrst 6 months of life is likely to be monogenic and not type 1 diabetes. This is supported by studies of monogenic causes of neonatal diabetes [1] and HLA studies [2,3]. Neonatal diabetes can be transient (if it remits), or permanent (lifelong diabetes) [4,5].

Transient Neonatal Diabetes (TNDM) Approximately half of patients with neonatal diabetes have TNDM [4] (OMIM#601410). More than 70% have abnormalities of an imprinted region on chromosome 6q24; paternal isodisomy, paternal duplication or methylation defects [5,6]. Transient neonatal diabetes is diagnosed at a median age of 3 days (0–31 days) [7]. These patients have a strikingly low birth (mean 1.98 ± 5.1 kg) [5] and at least 50% of patients will relapse and develop diabetes later in life [6]. In addition some patients with TNDM have mutations in the KCNJ11 or ABCC8 genes and they can also relapse after their initial period of remission [8,9].

Permanent Neonatal Diabetes (PNDM) There are more genetic etiologies of permanent neonatal diabetes (OMIM#606176) than TNDM. Most of the genes are either transcription factors involved in beta-cell development or critical for the function of the mature pancreatic beta cell (Fig. 1A).

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A

Hnf6 Ipf1* Ptf1a* Nkx2.2 Nkx6.1

Pluripotent precursor cell within the dorsal and ventral pancreatic buds e9.5-10

Hnf1beta* Ipf1* Precursor ductal cell e13-17

Ptf1a* Hes1

Ipf1* Ngn3 Ngn3 cell e14-17 Neurod1* Isl1 Pax4 Pax6 Nkx2.2 Nkx6.1 Hnf1a*

Endocrine pancreas beta cell e18-19

Precursor acinar cell e12.5-17

Exocrine pancreas acinar cell e18-19

Fig 1. A A schematic of the expression patterns of transcription factors during pancreatic development. The transcription factors required for each stage of development are shown irrespective of the order of expression. The expression patterns are based on genetic knock out studies in vivo or in vitro, adapted from Maestro et al. [21] and Servitja and Ferrer [139]. The transcription factors associated with diabetes are in bold and are marked with an asterisk

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B

Glucose GLUT2 transporter

Nucleus

Glyc

HNF1A* HNF4A* NEUROD1* IPF1* HNF1beta* Ins uli

Glucose

nt ran

sc ri p tio n

olys

is

GCK*

ATP

ATP ATP

Ca2+ Insulin release

Ca2+

KATP Channel (Kir6.2* & SUR1* subunits) K+

Ca2+ Ca2+

K+ K+

Ca2+ Channel o Dep

lari

sat

ion

Fig 1. B The overall function of the mature pancreatic beta cell, illustrating glucose mediated insulin secretion. The mechanism is triggered by glucose entering the cell through the GLUT2 transporter. Once inside the beta-cell, glucose is phosphorylated by glucokinase to glucose-6-phosphate, the substrate of glycolysis. Metabolism of glucose increases the level of ATP within the cell. ATP closes the inwardly rectifying potassium channel (KATP), causing depolarization of the cell membrane. In response to changes in cell membrane potential the voltage sensitive Ca2+ channel opens. Ca2+ inﬂux initiates insulin exocytosis, this requires an increase in transcription of the insulin gene via the pancreatic transcription factors. The genes associated with monogenic diabetes are essential to the mechanism of insulin release in the mature beta cell, these are highlighted in blue and marked with an asterisk

Altered Transcription Factor Function in Beta-Cell Development Mutations in the transcription factors involved in the development of the pancreas and the maturation of beta cells can present as neonatal diabetes.

Mutations in IPF1 Molecular Genetics There are two families described to date with a homozygous single base pair deletion or compound heterozygous missense IPF1 mutations [10,11].

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Clinical Phenotype Both patients had severe pancreatic agenesis, with an absent pancreas on CT scan [10,11] (OMIM#260370). They had marked intrauterine growth retardation (<10th centile). Heterozygous mutations result in early–onset diabetes (MODY4) [12] (discussed in section on IPF1 MODY below). Mechanism of Disease IPF1 (also known as PDX1) is a homeodomain transcriptional regulator required for pancreatic and beta-cell development. The pancreatic agenesis seen in these patients reﬂects this role and mirrors the pancreatic agenesis seen in knockout mice [13].

Mutations in PTF1A Molecular Genetics There have been 3 patients from 2 consanguineous families reported with homozygous PTF1A mutations [14]. Clinical Phenotype Neonatal diabetes is diagnosed on the ﬁrst day of life with severe hyperglycemia requiring insulin treatment due to pancreatic agenesis (OMIM#260370). Additional features include severe intrauterine growth retardation and severe neurological dysfunction with cerebellar hypoplasia, dysmorphic facial features, limb stiffness, and limb contractures [14,15]. Mechanism of Disease PTF1A is essential for pancreatic development and the maintenance of the adult exocrine pancreas [16,17] (Fig. 1A). PTF1A is also expressed in the developing neural tissue of the mouse, with expression in the cerebellum extending later in embryonic development [18]. Ptf1a knockout mice have cerebellar agenesis at birth conﬁrming the importance of the gene in neurogenesis as well as in pancreatic development [14].

Mutations in Hepatocyte Nuclear Factor-1beta (HNF-1beta) Molecular Genetics There have been two patients with neonatal diabetes who have different heterozygous HNF-1beta (also known as TCF2) mutations affecting amino acid S148 [19,20]. This is an unusual phenotype for HNF-1beta mutations which usually cause diabetes later in life and result in abnormalities of renal development (discussed later in section on HNF1beta MODY). Clinical Phenotype Both patients showed remission of diabetes with later relapse. They also had disorders of renal development which led to the investigation of HNF-1beta.

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Interestingly, other patients with the same mutations were not diagnosed with neonatal diabetes suggesting phenotypic variation [19,20]. Mechanism of Disease Hepatocyte nuclear factor-1beta is a transcription factor that functions either as a homodimer or heterodimer with HNF-1alpha. HNF-1beta is an important and early-acting member of the transcriptional network that regulates the generation of pancreatic progenitor cells that become both endocrine and exocrine pancreatic cells in the mouse [21,22] (Fig. 1A). Therefore patients with HNF-1beta mutations have both exocrine and endocrine pancreatic dysfunction. In keeping with this key developmental role, patients with a HNF-1beta mutation have a low birth weight (due to reduced insulin secretion in utero), exocrine dysfunction and reduced pancreatic size [20].

Mutations in GLIS3 Molecular Genetics Homozygous mutations in GLIS3 have been reported in 4 patients with neonatal diabetes from three consanguineous families [23]. Clinical Phenotype Mutations in GLIS3 are associated with neonatal diabetes and congenital hypothyroidism (the NDH syndrome) (OMIM#610199) [23]. At birth all patients have severe intrauterine growth retardation and neonatal diabetes is diagnosed soon afterwards. Other clinical features include facial dysmorphology, congenital glaucoma, liver ﬁbrosis, cystic kidneys, mild mental retardation, and often early infant death [23,24]. Patients with a GLIS3 mutation do not have the exocrine insufﬁciency found in patients with mutations in other pancreatic transcription factors. Mechanism of Disease The GLIS3 gene encodes GLI similar 3, a transcription factor involved in pancreatic development and pancreatic beta cells. The gene has multiple transcripts, which are expressed in a variety of tissues, such as liver, kidney, skeletal muscle, heart, kidney, thyroid, and pancreas [23]. This expression proﬁle is consistent with the multi-system syndrome associated with loss of function GLIS3 mutations in man.

Beta-Cell Destruction Early and severe beta-cell destruction can result in neonatal diabetes.

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Mutations in Eukaryotic Translation Initiation Factor-2 A Kinase 3 (EIF2AK3) Molecular Genetics Eighteen children, mostly from consanguineous families, have been described with Wolcott–Rallison syndrome (OMIM#226980) as a result of homozygous or compound heterozygous mutations in the eukaryotic translation initiation factor-2 alpha kinase 3 (EIF2AK3;PERK) gene [25]. Clinical Phenotype Diabetes is diagnosed in Wolcott–Rallison syndrome at a median of 13 weeks (range 6–65), with the majority requiring constant insulin treatment. Errors in bone development including multiple epiphyseal dysplasia, short stature, osteoporosis, or fractures are common. Other manifestations include: enlarged liver, liver disease which commonly leads to liver failure, developmental delay, renal failure, pancreatic hypoplasia, and early death [25–29]. Mechanism of Disease The enzyme EIF2AK3 phosphorylates EIF2alpha at serine 51. EIF2alpha phosphorylation regulates the synthesis of unfolded proteins in the endoplasmic reticulum (ER) and is enriched in pancreatic cells [30,31]. EIF2AK3 is expressed in developing pancreas, bone, kidney, adult liver, and adult beta cells as well as in exocrine tissue. Mutations in the EIF2AK3 gene abolish the enzyme’s kinase activity and ability to phosphorylate its target EIF2alpha [26]. This results in an increased number of unfolded proteins in the ER causing ER stress, which has been shown to cause beta-cell apoptosis in mice [26].

Mutations in the Fork Head Box Protein P3 (FOXP3) Molecular Genetics Mutations in Fork Head Box P3 (FOXP3) result in the immunodysregulation, polyendocrinopathy, and enteropathy, X-linked (IPEX) syndrome (OMIM#304790) [32]. Nineteen cases have been described to date and all are males as it is an X-linked recessive disease. Clinical Phenotype IPEX syndrome usually presents with diarrhea, autoimmune enteropathy, and/or autoimmune diabetes in the ﬁrst months of life [33]. Other clinical manifestations include anemia, hypothyroidism, recurrent infections, eczema, and atopic dermatitis. Pancreatic and thyroid auto-antibodies may be detected. Infantile death can occur from hemorrhage, diabetic complications, sepsis, diarrhea, or malnutrition Treatment with immunosuppression and bone marrow transplantation has varying degrees of success [34]. Female mutation carriers are healthy.

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Mechanism of Disease FOXP3 encodes the scurﬁn protein, which appears to act as a differentiation switch for the development of T regulatory cells. Therefore, a mutation will disrupt the immune system resulting in a loss of beta-cell mass and immune destruction of intestinal cells. The mouse model identiﬁes the underlying defect in IPEX as a lack of CD4+CD25+ T regulatory cells [35].

Defects in Beta-Cell Function Recently it has been shown that the most common cause of neonatal diabetes are defects in insulin secretion within the mature beta cell.

Activating Mutations in the Inwardly Rectifying Subunit, Kir6.2 (KCNJ11) of the KATP Channel Molecular Genetics The KCNJ11 gene encodes the inwardly rectifying subunit (Kir6.2) of the ATP sensitive potassium channel. Heterozygous activating mutations in KCNJ11 are the most common cause of PNDM (found in ∼112 families) in multiple populations and ethnic groups (OMIM#606176) [36]. Mutations are located throughout the single exon gene but there are two mutation hotspots, amino acids V59 and R201, where there are CpG dinucleotides [1]. KCNJ11 mutations are usually spontaneous but in 20% of cases there is autosomal dominant inheritance. Germline mosaicism has also been described [37]. Clinical Phenotype All KCNJ11 mutation carriers have marked hyperglycemia and are diagnosed with diabetes in the ﬁrst 6 months of life, median age 5 weeks (range 0–26), and in many cases present with ketoacidosis [1]. They have a low birth weight (<10th centile) due to reduced insulin secretion in utero [38]. Thirty percent of patients have neurological features. The most severe neurological phenotype is known as the Developmental delay, Epilepsy, and Neonatal Diabetes (DEND) syndrome. These children are unable to walk or talk and have a poor prognosis [39]. There is also a milder and more common subtype known as intermediate DEND (I-DEND), where mutation carriers have moderate developmental delay and muscle weakness but do not have epilepsy. This is usually associated with the V59M mutation [1,39]. Identifying the genetic etiology of neonatal diabetes has important implications for treatment. Patients with isolated diabetes and I-DEND can usually transfer off insulin and onto sulfonylurea tablets, and achieve excellent glycemic control [40–42]. Long-term follow-up studies are needed but initial studies suggest this treatment response may be long lasting [42]. Patients with DEND syndrome do not respond to sulfonylureas in keeping with in vitro

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studies showing these mutations show a reduced response to sulfonylureas in vitro [43,44]. Mechanism of Disease The role of the Kir6.2 subunit is to bind intracellular ATP, produced from glycolysis within the beta cell, and result in KATP channel closure and hence depolarization of the cell membrane and insulin release (Fig. 1B). Gain of function KCNJ11 mutations reduce the ability of ATP to block the KATP channel or increase the probability of the channel being in the open state, depending on the location of the mutations [36,45]. The gene is expressed in the pancreatic beta cell, smooth muscle, brain, heart and neurons, therefore the observed phenotypes reﬂect the altered KATP channel activity in these tissues. Sulfonylurea drugs close the KATP channel through an ATP independent route by binding to the SUR1 subunit of the channel. The functional properties of the KATP channel are described in the chapter by Proks and Ashcroft in this volume.

Activating Mutations in the Sulfonylurea Receptor-1, SUR1 (ABCC8) Subunit of the KATP Channel Molecular Genetics Gain of function heterozygous mutations in the ABCC8 gene, which encodes the sulfonylurea receptor-1 (SUR1), can also cause PNDM. Three cases have recently been reported [9,46], and nine additional cases have been identiﬁed in our laboratory (unpublished data from S.E. Flanagan, A.-M. Patch, S. Ellard, and A.T. Hattersley). Mutations in ABCC8 have also been identiﬁed in 20 patients with TNDM ([9] and unpublished data from S.E. Flanagan, A.-M. Patch, S. Ellard, and A. T. Hattersley). Phenotypic Features Heterozygous activating mutations in ABCC8 result in diabetes diagnosed in the ﬁrst few months of life. A single case had marked neurological features including motor and social developmental delay, muscle weakness and epilepsy, as seen in the DEND syndrome associated with Kir6.2 mutations [46]. Mechanism of Disease The KATP channel is an octameric protein composed of four SUR1 and Kir6.2 subunits. KATP channel closure initiates depolarization of the beta-cell membrane. The channel opens when magnesium nucleotides bind to SUR1 whereas channel closure is controlled by nucleotide binding to Kir6.2. Heterozygous activating mutations in SUR1 increase the open probability of the channel due by increasing the afﬁnity for magnesium nucleotides [9] or by making it less sensitive to ATP[46]. Identifying a ABCC8 mutation may have treatment

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advantages as sulfonylureas can replace insulin treatment in some patients ([9] and unpublished data S.E. Flanagan, S. Ellard, and A.T. Hattersley).

Inactivating Glucokinase Mutations Molecular Genetics Six patients have been reported with neonatal diabetes caused by homozygous or compound heterozygous mutations in the gene encoding the glycolytic enzyme glucokinase (GCK) [47] (OMIM#606176). Heterozygous inactivating mutations cause mild, non-progressive, hyperglycemia (discussed in section on GCK MODY below), whereas heterozygous activating mutations cause familial hyperinsulinemic hypoglycemia [48]. Clinical Phenotype Permanent neonatal diabetes arising from glucokinase mutations presents in the ﬁrst few days of life and requires insulin treatment. All cases have marked intrauterine growth retardation. Parents with heterozygous GCK mutations have glucose intolerance with fasting hyperglycemia >5.5 mmol/l. Mechanism of Disease The glucokinase enzyme phosphorylates glucose and is predominantly expressed in the beta cell and hepatocyte. In the pancreatic beta cell glucokinase is the “glucose sensor” linked to insulin secretion (Fig. 1B). Homozygous inactivating GCK mutations result in a complete loss of the enzyme activity, therefore glucose phosphorylation and insulin secretion is abolished, resulting in permanent neonatal diabetes [47,49]. The role of glucokinase in the regulation of insulin secretion is described in more detail in the chapter by Matschinsky in this volume.

Permanent Neonatal Diabetes of Unknown Cause The eight known genetic causes of PNDM, detailed above, do not explain all patients with neonatal diabetes; ∼20% of patients have an unknown genetic etiology (which we term PNDM X) (Fig. 2A). Some may be atypical cases, for example mutations in the glucose transporter GLUT2 cause Fanconi–Bickel syndrome [50], neonatal diabetes is not a feature of this but a single case with neonatal diabetes has been described [51]. The future of determining the molecular basis of PNDM X is in the hands of geneticists who will use both a candidate gene approach (possibly targeting other transcription factor genes involved in beta-cell development, see Fig. 1A) and in linkage studies large consanguineous families to deﬁne new causes of neonatal diabetes.

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A Unknown aetiology

PNDM

Defects in pancreatic Defects in beta-cell development function

KCNJ11 ABCC8 40% 15%

PNDM X 20%

Beta-cell destruction

GCK 2%

EIF2AK3 10%

FOXP3 10%

Pancreatic beta-cell transcription factors GLIS3 2%

HNF-1beta <1%

IPF1 <1% MODY

B

Unknown aetiology

Defects within the beta cell Defects within the exocrine pancreas GCK

CEL <1%

22%

MODY X 11%

Pancreatic beta-cell transcription factors HNF1A

HNF4A

58%

5%

HNF-1beta 2%

IPF1

NEUROD1

<1%

<1%

Fig. 2A,B. Diagram showing the distribution of known genetic causes of permanent neonatal diabetes and maturity-onset diabetes of the young, adapted from Hattersley and Pearson [140]

Genetics of PNDM: Conclusion In the past 5 years there have been considerable advances in deﬁning the molecular genetics of PNDM. These have not only helped reclassify patients and give scientiﬁc insights into the beta cell but it has also helped patient care. The transfer of children and adults with KCNJ11 mutations from insulin to sulfonylureas has improved glycemic control as well as improving quality of

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life. The clinical characteristics of patients can be used to help determine which molecular genetic test should be carried out ﬁrst (Table 1). There are still a group of patients without a known genetic etiology, and these patients are likely to have mutations in critical beta-cell genes.

Maturity-Onset Diabetes of the Young: The Clinical Description Maturity-onset diabetes of the young (MODY) was initially deﬁned as autosomal dominant inheritance of non-insulin dependent diabetes, diagnosed before 25 years of age (OMIM#606391) [52, 53]. Our understanding of MODY has been transformed by the identiﬁcation of new genetic etiologies. These novel ﬁndings have furthered our understanding of the underlying disease mechanism of beta-cell dysfunction. MODY now encompass a heterogeneous group of disorders, with each being a discrete clinical and physiological entity (Table 2). This work has advanced our understanding of early onset diabetes and has led to diagnostic genetic testing which can determine the likely clinical course, prognosis and best treatment options for patients.

Errors in Glycolytic Enzymes Glucokinase (GCK MODY) Molecular Genetics Glucokinase (GCK) was the ﬁrst MODY gene to be identiﬁed [54,55] (OMIM#125851). Over 200 mutations in GCK are distributed throughout the gene and these include nonsense, missense, frameshift and splice site mutations [56]. Homozygous or compound heterozygous mutations result in permanent neonatal diabetes (discussed in section on Inactivating Glucokinase Mutations above). Clinical Phenotype Mutations in GCK result in a discrete phenotype despite the wide variety of mutations [57]. In the UK 30% of monogenic diabetes is caused by a GCK mutation [58]. Mutations result in a mild stable fasting hyperglycemia with slow deterioration of beta-cell function over time (as seen in the normal population). A fasting glucose below 5.5 mmol/l is extremely rare in GCK MODY patients, and the 0–2 h oral glucose tolerance test (OGTT) increment is below 3 mmol/l in 71% of mutation carriers [59,60]. In contrast to transcription factor MODY, mild hyperglycemia is present from birth, and patients are often asymptomatic [61].

2.58 kg (−1.73)

≅100 Rare

18 90%

32 Rare

PNDM TNDM

PNDM

PNDM TNDM

EIF2AK3 (Wolcott– Rallison syndrome) Recessive

ABCC8 Spontaneous & autosomal dominant

3.05 kg (−1.05)

?

2.10 kg (−2.94)

ZAC/HYAMI imprinting defect on 6q24 KCNJ11 Spontaneous (∼80%) & autosomal dominant

≅150 Rare

% in consanguineous or isolated populations

TNDM

Inheritance

Table 1. Genetic and clinical characteristics of diabetes in the ﬁrst 6 months of life Gene PNDM/TNDM No. of families Median birth described with a weight (SDS) mutation

4 (0.5–18)

13 (6–65)

6 (0–260)

0.5 (0–4)

Age of diagnosis in weeks. Median (range)

Normal

Exocrine dysfunction (25%)

Normal

Normal

Pancreatic appearance

Developmental delay (20%), epilepsy (6%), DKA (30%), sulfonylurea treated Epiphyseal dysplasia (90%), osteopenia (50%), acute liver failure (75%), developmental delay (80%), hypothyroidism (25%) Developmental delay, epilepsy, sulfonylurea treated

Macroglossia (23%)

Other features

Genetic Disorders of the Pancreatic Beta Cell and Diabetes 411

19 Rare

6 85% 2 50%

PNDM

PNDM

GCK Recessive

IPF1 Recessive

% in consanguineous or isolated populations

No. of families described with a mutation

PNDM

PNDM/TNDM

FOXP3 (IPEX syndrome) X linked

Inheritance

Table 1. Continued Gene

2.14 kg (−3.0)

1.72 kg (−2.8)

2.86 kg (−1.2)

Median birth weight (SDS)

–

0.14 (0.14–1.6)

6 (0–30)

Age of diagnosis in weeks. Median (range)

Absent

Normal

?

Pancreatic appearance

Only boys affected. Chronic diarrhea (95%), pancreatic and thyroid autoantibodies (75%), thyroiditis (20%), eczema (50%), anemia (30%), often die young (1st year) Parents have fasting hyperglycemia as heterozygotes Parents have earlyonset diabetes as heterozygotes

Other features

412 E.L. Edghill and A.T. Hattersley

2 Rare 2 100%

3 100%

TNDM

PNDM

PNDM

DKA, Diabetic ketoacidosis; SDS, standard deviation score

GLIS3 Recessive

TCF2/HNF1beta Spontaneous & autosomal dominant PTF1A Recessive

1.79 kg (−3.9)

1.39 kg (−3.8)

1.90 kg (−3.2)

0.28 (0.14–0.29)

0.14

2.28 (2.14–2.43)

?

Atrophy

Atrophy Renal disease (e.g. renal cysts, renal failure) Severe neurological dysfunction, cerebellar hypoplasia, ﬂexion contractures Congenital hypothyroidism, facial dysmorphology, congenital glaucoma, cystic kidneys, liver ﬁbrosis, early death

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Adolescence and early adulthood Early adulthood Early adulthood

Early adulthood Adulthood Rarely in early adulthood

>95% >80% >95%

>80% >80% >99%

49%

<1%

6%

<1% <1%

<1% diagnosed in early adulthood (97%? diagnosed in ﬁrst 6 months of life)

IPF1

TCF2/HNF-1beta

NEUROD1 CEL

KCNJ11/ABCC8

a

Data from Gloyn and Ellard [58]

TCF1/HNF1A

Early childhood

>45%

31%

GCK

Adolescence and early adulthood

10%

HNF4A

>90%

Table 2. Characteristics of diabetes presenting in childhood or early adulthood Age of onset Frequency in the UK Gene Penetrance of mutations by age monogenic diabetes 30 years populationa

Limited data

Limited data Limited data

Frequent due to progressive hyperglycemia

Rare due to mild hyperglycemia Frequent due to progressive hyperglycemia Limited data

Frequent due to progressive hyperglycemia

Microvascular complications

Pancreatic agenesis in homozygotes Renal disease, abnormal liver tests, genital tract abnormalities, reduced birth weight, pancreatic atrophy Limited data Pancreatic atrophy, loose stool —

Sensitive to sulfonylureas

Transient hypoglycemia at birth, high birth weight, low triglycerides Reduced birth weight

Other features

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GCK is important in glucose sensing in utero and babies with a GCK mutation have a reduced birth weight as a result of reduced insulin secretion in utero [62]. In most patients the mild hyperglycemia is detected by screening either during routine medicals in pregnancy, or family screening when MODY is suspected [63]. In contrast to transcription factor MODY, patients with glucokinase mutations rarely need any pharmacological treatment [60,64,65]. The majority (>85%) are managed on diet alone [59,60]. Mechanism of Disease Glucokinase is principally expressed in pancreatic beta cells and in hepatocytes. In beta cells GCK can change the rate of glucose phosphorylation depending on the physiological concentration of glucose, and hence it closely links insulin secretion with the glucose concentration [66] (Fig. 1B). Mutations alter either the enzyme’s maximal activity for glucose phosphorylation, the afﬁnity for glucose [67], causing a major alteration in protein structure [68], or protein expression. These reductions in function result in reduced activity and hence a resetting of the glucose concentration to a higher fasting value.

Errors in Pancreatic Beta-Cell Transcription Factors The MODY transcription factors form a regulatory network which maintains beta-cell function and hence normal glucose homeostasis [69] (Fig. 1B). HNF1alpha and HNF1beta are homeodomain-containing transcription factors. They share 90% amino acid homology in their DNA binding domain and they dimerize to form homo- or heterodimers; they also recognize the same DNA binding site [70]. HNF-1alpha interacts with many of the other transcription factors, with a transcriptional activation feedback loop existing between HNF-1alpha and HNF-4alpha [71,72].

Hepatocyte Nuclear Factor-4alpha (HNF4A MODY) Molecular Genetics The MODY1 locus was the ﬁrst MODY gene to be identiﬁed by linkage studies [73]. Subsequently the gene was identiﬁed as HNF4A [74]. Mutations in HNF4A are considerably less common than TCF1 (HNF-1alpha gene) [75], and in the UK they account for 10% of cases with monogenic diabetes [58]. Mutations are distributed throughout the coding region and pancreatic (P2) promoter [76]. Clinical Phenotype Mutations in HNF4A result in MODY (OMIM#125850). The phenotype is very similar to that seen in TCF1 mutation carriers although the age of diagnosis

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is slightly later [77]. Patients with HNF4A mutations have progressive betacell dysfunction, characterized by an inability to increase insulin secretion at high blood glucose levels [78]. Some patients are successfully treated with sulfonylureas [79] and these patients are sensitive to the hypoglycemic effects of sulfonylurea therapy. In contrast to HNF1A mutation carriers, that have normal birth weight, neonates are frequently macrosomic and may present with hyperinsulinemic hypoglycemia [80]. Another distinguishing feature is the altered apolipoprotein and triglyceride concentrations [77,81]. Disease Mechanism Mutations in HNF4A result in haploinsufﬁciency (loss of function/gene dosage effect) and hence there is a reduction in target gene transcription [75]. There is a positive feedback loop existing between HNF-1alpha and the pancreatic (P2) promoter of HNF4A which may act as a switch explaining some similarities in the beta-cell phenotype between HNF1A and HNF4A mutation carriers [71,72]. This also explains how mutations in the pancreatic P2 promoter cause MODY [82, 83]. The mechanism for the transient hypoglycemia which appears to be unique to HNF4A is uncertain. A beta-cell speciﬁc HNF4A knock out mouse also shows hypoglycemia, and it was suggested that this resulted from reduced KCNJ11 transcription and hence decreased expression of this KATP channel subunit [84].

Hepatocyte Nuclear Factor-1alpha Mutations (HNF1A MODY) Molecular Genetics Hepatocyte nuclear factor-1alpha (HNF1A: TCF1) mutations are the most common cause of MODY (OMIM#600496) [85,86]. Over 200 mutations have been described arising throughout the promoter, coding region, and conserved splice sites [76]. The most commonly reported mutation is Pro291fsinsC (17% of families) that occurs in a poly C tract which are prone to deletion or duplication of a base by slipped mispairing at the replication fork [76]. Clinical Phenotype The median age of diagnosis of diabetes is 21 years and the majority of patients are diagnosed before 30 years although a few individuals can reach middle age without developing diabetes [87]. Interestingly the position of the mutation relative to the different beta-cell isomers of HNF-1alpha can impact on the age of diagnosis; mutations which compromise all three isoforms are diagnosed on average 7 years earlier than those which affect only single isomers or groups of isoforms [88]. In childhood or adolescence, subjects with HNF1A mutations usually have normal glucose tolerance [42]. An early diagnosis of diabetes can be made as a result of an oral glucose tolerance test since fasting blood glucose may initially be normal [59]. Prior to developing diabetes HNF1A mutation carriers have marked beta-cell deﬁciency, increased

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insulin sensitivity, and glycosuria [89]. Glycosuria is an early clinical marker due to the reduced renal threshold seen in these patients [89–91]. HNF1A mutation carriers have a progressive beta-cell defect [60,92,93] leading to a failure to increase insulin secretion appropriately in response to hyperglycemia. Microvascular and macrovascular complications are common making it important that they achieve good glycemic control [94,95]. A diagnosis of HNF1A MODY alters the choice of therapy. Mutation carriers show marked sensitivity to the hypoglycemic action of sulfonylureas compared to type 2 diabetes patients [96–98]. This may result in marked symptomatic hypoglycemia while starting therapy, but low-dose sulfonylureas are the appropriate ﬁrst-line medication for patients with HNF-1alpha mutations [97]. Disease Mechanisms Like HNF4A MODY, HNF1A MODY is also a disease of haploinsufﬁciency. Although initial studies suggested that the commonest mutation Pro291fsinsC acted as a dominant negative mutation [99], subsequent studies have shown that the mutant mRNA transcript is degraded by nonsense-mediated decay and hence a dominant negative-acting protein is unlikely to be produced in vivo [100]. Reduced HNF-1alpha activity in the beta cell results in reduction of many critical components of carbohydrate metabolism, including reduced GLUT2 (a glucose transporter) expression and mitochondrial dysfunction [101–103]. These early defects in metabolism probably explain the sensitivity to sulfonylureas which act downstream of these defects [98]. Mutations may also result in progressive beta-cell deﬁciency by increased beta-cell apoptosis and reduced beta-cell proliferation [103,104]. The observed low renal threshold resulting in glycosuria may be due to reduced expression of the high capacity/low afﬁnity sodium-glucose transporter-2 (SGLT-2) reducing glucose reabsorption in the proximal tubule [105].

Mutations in Insulin Promoter Factor 1 (IPF1 MODY) Molecular Genetics Heterozygous mutations in the insulin promoter factor 1 (IPF1) gene is a very rare cause of MODY (OMIM#606392). Homozygous mutations result in permanent neonatal diabetes and pancreatic agenesis (discussed in section on IPF1 Mutations above). Clinical Phenotype The clinical phenotype of heterozygous IPF1 mutation carriers is similar to other types of transcription factor MODY but they tend to present later [12,106].

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Disease Mechanism IPF1 is a transcription factor crucial for the early stages of pancreatic and beta-cell development and is also involved in the transcription of key beta-cell genes. Heterozygous knockout in mice causes impaired glucose tolerance and have a reduction in transcription of beta-cell speciﬁc genes such as insulin, Nkx6.1 and Glut2 [107], which is probably at least part of the mechanism that results in IPF1 mutations causing MODY in man.

Hepatocyte Nuclear Factor-1beta Mutations (Renal Cysts and Diabetes Syndrome/HNF1beta MODY) Molecular Genetics Heterozygous mutations in the gene encoding HNF-1beta (TCF2) were identiﬁed as a cause of MODY soon after the description of HNF1A mutations [108]. This is usually an autosomal dominant disease, but spontaneous mutations are common. Mutations include missense, nonsense, splice site, insertions and deletions [109,110]. Heterozygous gross deletions on chromosome 17 encompassing the HNF1beta gene account for approximately a third of the known mutations ([111,112] and unpublished data from E.L. Edghill, A.T. Hattersley, S. Ellard, and C. Bingham). Clinical Phenotype The HNF1beta mutation phenotype is strikingly different to the other MODY transcription factor phenotypes and rarely presents just as diabetes [110]. The primary phenotype is nondiabetic renal disease, in particular renal cysts, with presentation antenatally or early in life. Early-onset diabetes is observed in most cases, but this presents later than the renal disease (∼20 years of age) [110]. The diabetes unlike other forms of MODY is associated with increased insulin resistance and rapidly requires treatment with insulin [113]. Renal Cysts And Diabetes (RCAD) is used to describe the commonest clinical syndrome associated with HNF1beta mutations (OMIM#137920) [109]. The main renal phenotypes include cysts [114], glomerulocystic kidney disease [115], and structural kidney disease [116]. Other clinical features include genital tract abnormalities [117], pancreatic atrophy [118], neonatal diabetes [19], liver dysfunction [119], and biliary dysfunction [120]. There is no difference in phenotype between those with a mutation and a whole gene deletion [111,112]. Patients frequently have a low birth weight (<10th centile) [20]. Disease Mechanism HNF1beta mutations in man probably result in diabetes by a reduction in beta-cell development consistent with their reduced insulin-mediated fetal growth and progressive beta-cell defect postnatally. The role of HNF-1beta in the development of the pancreas has been difﬁcult to study in knockout

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animals as HNF1beta-null mice die at the blastocyst stage (e7.5d) [121,122]. HNF1beta-null embryos, rescued from early lethality by using embryonic stem cells, had pancreatic agenesis by embryonic day 13.5, showing the key role of HNF-1beta in both endocrine and exocrine murine pancreatic development [22]. Elegant studies by Maestro and colleagues have shown that cells expressing HNF1beta are the immediate precursors of Neurogenin3-positive cells, the progenitors from which endocrine cells arise during pancreatic organogenesis [21] (Fig. 1A). HNF-1beta is also a regulator of HNF6 expression, another transcription factor involved in pancreas development [123]. The variety of renal manifestations observed in HNF1beta mutation carriers highlights its importance in kidney development where it is expressed very early in development. Renal cysts are the most common renal phenotype. This may occur through the disruption of Pkhd1 and Umod expression, two of HNF-1beta’s transcriptional targets [124]. Pkhd1 and Umod mutations cause autosomal-recessive polycystic kidney disease [125] and familial juvenile hyperuricemic nephropathy [126], respectively.

Mutations in NEUROD1 (NEUROD1 MODY) Molecular Genetics Mutations in NEUROD1 have been described in three families but in only one is there complete co-segregation with diabetes (OMIM#606394) [127]. Clinical Phenotype Diabetes presents over a wide age range and some cases have long term complications [127,128]. Disease Mechanism NEUROD1 is a basic helix loop helix (HLH) transcription factor. It functions as a regulatory switch for the endocrine pancreas where it controls the expression of the insulin gene [129]. NeuroD1-null mice have abnormal islet development and on some genetic backgrounds diabetes as well [130]. NEUROD1 is a transcriptional activator, and functions through binding to the coactivators CBP and p300 [131,132]. Mutations within the binding domain can reduce binding to co-activators and target genes.

Defects in Exocrine Enzymes Mutations in Carboxy Ester Lipase (CEL MODY) Molecular Genetics Mutations in the tandem repeats (VNTR), located in exon 11 of the carboxy ester lipase (CEL) gene were recently shown to cause diabetes in a families that had MODY by clinical criteria [133].

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Clinical Phenotype Patients with a single base deletion in the CEL VNTR had diabetes, presenting at 34 ± 12 years, and requiring insulin treatment. Preceding diabetes, by the age of 10 years, patients have fecal elastase deﬁciency, mild abdominal pain, and loose stools, an indicator of exocrine dysfunction. Most patients have pancreatic atrophy reﬂecting their endocrine and exocrine pancreatic dysfunction [133]. Disease Mechanism Carboxyl ester lipase is expressed in acinar cells, not in beta cells [134]. It is an important component of the pancreatic juice responsible for breaking down cholesterol esters in adults and milk fat in neonates [134,135]. It is uncertain at present how the primary exocrine dysfunction results in beta-cell dysfunction.

MODY X Families There are a signiﬁcant proportion of MODY families where the genetic etiology remains unknown. In the UK this accounts for about 11% of strictly deﬁned MODY families [136] (Fig. 2B). These MODY X families ﬁt classical MODY criteria but do not have mutations in any of the known MODY genes. Key candidate genes are likely to be involved in gene transcription, pancreatic development, or glucose sensing in the pancreatic islet. However, there is evidence that this may not be the only place to explore; as genetic mutations affecting the exocrine pancreas can cause beta-cell dysfunction and diabetes [133,137].

MODY: Conclusion Mutations in seven discrete genes have been shown to cause MODY: GCK, TCF1, TCF2, HNF4A, IPF1, NEUROD1 and CEL. The description of these genes has allowed us to understand much of the clinical and physiological heterogeneity observed in MODY [138]. Molecular genetic testing will beneﬁt patients and clinicians by being able to make a speciﬁc diagnosis and guiding clinical management. Acknowledgments. We are very grateful to Diabetes UK, the Wellcome Trust, DIRECT, The Royal Devon and Exeter NHS Foundation Trust, Peninsula Medical School, and the University of Exeter for supporting our work on monogenic diabetes. We would like to acknowledge all our colleagues at Exeter and our collaborators throughout the world who have contributed to the ideas in this chapter.

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92. Byrne MM, Sturis J, Menzel S, Yamagata K, Fajans SS, Dronsﬁeld MJ, Bain SC, Hattersley AT, Velho G, Froguel P, Bell GI, Polonsky KS (1996) Altered insulin secretory responses to glucose in diabetic and nondiabetic subjects with mutations in the diabetes susceptibility gene MODY3 on Chromosome 12. Diabetes 45:1503–1510 93. Hattersley AT (1998) Maturity-onset diabetes of the young: Clinical heterogeneity explained by genetic hetergeneity. Diabet Med 15:15–24 94. Velho G, Vaxillaire M, Boccio V, Charpentier G, Froguel P (1996) Diabetes complications in NIDDM kindreds linked to the MODY3 locus on chromosome 12q. Diabetes Care 19:915–919 95. Isomaa B, Henricsson M, Lehto M, Forsblom C, Karanko S, Sarelin L, Haggblom M, Groop L (1998) Chronic diabetic complications in patients with MODY3 diabetes. Diabetologia 41:467–473 96. Sovik O, Njolstad P, Folling I, Sagen J, Cockburn BN, Bell GI (1998) Hyperexcitability to sulphonylurea in MODY3. Diabetologia 41:607–608 97. Pearson ER, Liddell WG, Shepherd M, Corrall RJ, Hattersley AT (2000) Sensitivity to sulphonylureas in patients with hepatocyte nuclear factor 1 alpha gene mutations: evidence for pharmacogenetics in diabetes. Diabet Med 17:543–545 98. Pearson ER, Starkey BJ, Powell RJ, Gribble FM, Clark PM, Hattersley AT (2003) Genetic aetiology of hyperglycaemia determines response to treatment in diabetes. Lancet 362:1275–1281 99. Yamagata K, Nammo T, Moriwaki M, Ihara A, Iizuka K, Yang Q, Satoh T, Li M, Uenaka R, Okita K, Iwahashi H, Zhu Q, Cao Y, Imagawa A, Tochino Y, Hanafusa T, Miyagawa J, Matsuzawa Y (2002) Overexpression of dominant-negative mutant hepatocyte nuclear fctor-1 alpha in pancreatic beta-cells causes abnormal islet architecture with decreased expression of E-cadherin, reduced beta-cell proliferation, and diabetes. Diabetes 51:114–123 100. Harries LW, Ellard S, Jones RW, Hattersley AT, Bingham C (2004) Abnormal splicing of hepatocyte nuclear factor-1 beta in the renal cysts and diabetes syndrome. Diabetologia 47:937–942 101. Wang H, Maechler P, Hagenfeldt KA, Wollheim CB (1998) Dominant-negative suppression of HNF-1alpha function results in defective insulin gene transcription and impaired metabolism-secretion coupling in a pancreatic beta-cell line. EMBO J 17:6701–6713 102. Wang H, Antinozzi PA, Hagenfeldt KA, Maechler P, Wollheim CB (2000) Molecular targets of a human HNF1 alpha mutation responsible for pancreatic beta-cell dysfunction. EMBO J 19:4257–4264 103. Shih DQ, Screenan S, Munoz KN, Philipson L, Pontoglio M, Yaniv M, Polonsky KS, Stoffel M (2001) Loss of HNF-1alpha function in mice leads to abnormal expression of genes involved in pancreatic islet development and metabolism. Diabetes 50:2472–2480 104. Wobser H, Dumann H, Kogel D, Wang H, Reimertz C, Wollheim CB, Byrne MM, Prehn JH (2001) Dominant-negative suppression of HNF-1alpha results in mitochondrial dysfunction, INS-1 cell apoptosis, and Increased sensitivity to ceramide-, but not to high glucose-induced cell death. J Biol Chem 27:27 105. Pontoglio M, Prie D, Cheret C, Doyen A, Lero C, Froguel P, Velho G, Yaniv M, Friedlander G (2000) HNF1 alpha controls renal glucose reabsorption in mouse and man. EMBO Rep 1:359–365 106. Gragnoli C, Stanojevic V, Gorini A, Von Preussenthal GM, Thomas MK, Habener JF (2005) IPF-1/MODY4 gene missense mutation in an Italian family with type 2 and gestational diabetes. Metabolism 54:983–988

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107. Ahlgren U, Jonsson J, Jonsson L, Simu K, Edlund H (1998) beta-cell-speciﬁc inactivation of the mouse Ipf1/Pdx1 gene results in loss of the beta-cell phenotype and maturity-onset diabetes. Genes Dev 12:1763–1768 108. Horikawa Y, Iwasaki N, Hara M, Furuta H, Hinokio Y, Cockburn B, Lindner T, Yamagata K, Ogata M, Tomonaga O, Kuroki H, Kasahar T, Iwamoto Y, Bell GI (1997) Mutation in hepatocyte nuclear factor-1b gene (TCF2) associated with MODY. Nat Genet 17:384–385 109. Bingham C, Hattersley AT (2004) Renal cysts and diabetes syndrome resulting from mutations in hepatocyte nuclear factor-1beta. Nephrol Dial Transplant 19:2703– 2708 110. Edghill EL, Bingham C, Ellard S, Hattersley AT (2006) Mutations in hepatocyte nuclear factor-1beta and their related phenotypes. J Med Genet 43:84–90 111. Bellanne-Chantelot C, Clauin S, Chauveau D, Collin P, Daumont M, Douillard C, Dubois-Laforgue D, Dusselier L, Gautier JF, Jadoul M, Laloi-Michelin M, Jacquesson L, Larger E, Louis J, Nicolino M, Subra JF, Wilhem JM, Young J, Velho G, Timsit J (2005) Large genomic rearrangements in the hepatocyte nuclear factor-1beta (TCF2) gene are the most frequent cause of maturity-onset diabetes of the young type 5. Diabetes 54:3126–3132 112. Ulinski T, Lescure S, Beauﬁls S, Guigonis V, Decramer S, Morin D, Clauin S, Deschenes G, Bouissou F, Bensman A, Bellanne-Chantelot C (2006) Renal phenotypes related to hepatocyte nuclear factor-1beta (TCF2) mutations in a pediatric cohort. J Am Soc Nephrol 17:497–503 113. Pearson ER, Badman MK, Lockwood CR, Clark PM, Ellard S, Bingham C, Hattersley AT (2004) Contrasting diabetes phenotypes associated with hepatocyte nuclear factor-1alpha and -1beta mutations. Diabetes Care 27:1102–1107 114. Nishigori H, Yamada S, Kohama T, Tomura H, Sho K, Horikawa Y, Bell GI, Takeuchi T, Takeda J (1998) Frameshift mutation, A263fsinsGG, in the hepatocyte nuclear factor-1 beta gene associated with diabetes and renal dysfunction. Diabetes 47:1354–1355 115. Bingham C, Bulman MP, Ellard S, Allen LI, Lipkin GW, Hoff WG, Woolf AS, Rizzoni G, Novelli G, Nicholls AJ, Hattersley AT (2001) Mutations in the hepatocyte nuclear factor-1beta gene are associated with familial hypoplastic glomerulocystic kidney disease. Am J Hum Genet 68:219–224 116. Carbone I, Cotellessa M, Barella C, Minetti C, Ghiggeri GM, Caridi G, Perfumo F, Lorini R (2002) A novel hepatocyte nuclear factor-1beta (MODY-5) gene mutation in an Italian family with renal dysfunctions and early-onset diabetes. Diabetologia 45:153–154 117. Lindner TH, Njolstad PR, Horikawa Y, Bostad L, Bell GI, Sovik O (1999) A novel syndrome of diabetes mellitus, renal dysfunction and genital malformation associated with a partial deletion of the pseudo-POU domain of hepatocyte nuclear factor1beta. Hum Mol Genet 8:2001–2008 118. Bellanne-Chantelot C, Chauveau D, Gautier JF, Dubois-Laforgue D, Clauin S, Beauﬁls S, Wilhelm JM, Boitard C, Noel LH, Velho G, Timsit J (2004) Clinical spectrum associated with hepatocyte nuclear factor-1beta mutations. Ann Intern Med 140:510–517 119. Montoli A, Colussi G, Massa O, Caccia R, Rizzoni G, Civati G, Barbetti F (2002) Renal cysts and diabetes syndrome linked to mutations of the hepatocyte nuclear factor-1 beta gene: description of a new family with associated liver involvement. Am J Kidney Dis 40:397–402 120. Kitanaka S, Miki Y, Hayashi Y, Igarashi T (2004) Promoter-speciﬁc repression of hepatocyte nuclear factor (HNF)-1 beta and HNF-1 alpha transcriptional activity by

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20. ATP-Sensitive Potassium Channels in Health and Disease Peter Proks and Frances M. Ashcroft

Summary. Since their discovery over 20 years ago, it has been recognized that adenosine triphosphate-sensitive potassium (KATP) channels play a critical role in insulin secretion. When these channels are open, insulin secretion is inhibited, and when they are shut, secretion is initiated. Consequently drugs, mutations, or changes in beta-cell metabolism that open KATP channels decrease insulin secretion and may cause diabetes, whereas those manipulations that close KATP channels have the opposite effect, increasing insulin secretion and hypoglycemia. This chapter reviews our current knowledge of the pancreatic beta-cell KATP channel, and discusses new data on its structure, structure–function relationships, and role in disease.

Introduction Adenosine triphosphate (ATP)-sensitive potassium (KATP) channels couple metabolism to the resting membrane potential and electrical activity of the cell. In some tissues, they also mediate the actions of hormones and transmitters. Their physiological role was ﬁrst elucidated in the pancreatic beta cell, where they were found to couple glucose metabolism to insulin secretion [1,2] (Fig. 1). At substimulatory glucose concentrations, K+ efﬂux through KATP channels produces a negative membrane potential that keeps voltage-gated Ca2+ channels closed (Fig. 1a). Because Ca2+ is needed for insulin granule release, this prevents insulin secretion. When plasma glucose levels increase, glucose uptake and metabolism lead to changes in the intracellular concentrations of adenine nucleotides that close KATP channels. Consequently the membrane depolarizes, which triggers Ca2+ channel opening, Ca2+ inﬂux, and insulin release (Fig. 1b). Sulfonylureas, drugs long established for type 2 Oxford Centre for Gene Function, Department of Physiology, University of Oxford, Parks Road, Oxford OX1 3PT, UK

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Fig. 1. Schematics of insulin secretion from the pancreatic beta cell. Insulin secretion is triggered by an increase in intracellular calcium ([Ca2+]i), which, in turn, is regulated by the electrical activity of the pancreatic beta cell. When metabolism is low (a), KATP channels are open, maintaining the membrane hyperpolarized and Ca2+ channels shut, so that [Ca2+]i remains low. When metabolism increases (b), KATP channels close, producing depolarization, electrical activity, and opening of voltage-gated Ca2+ channels. This leads to Ca2+ inﬂux and an increase in [Ca2+]i that triggers insulin release. Sulfonylureas block KATP channels directly, thereby triggering depolarization and insulin secretion independently of the metabolic state of the beta cell (c). Mutations in KATP channel genes that cause loss of function stimulate insulin secretion, producing congenital hyperinsulinism (d), whereas those that cause gain of function hyperpolarize the beta cell, preventing insulin release which leads to neonatal diabetes (e)

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diabetes therapy, stimulate insulin secretion by bypassing beta-cell metabolism and closing KATP channels directly (Fig. 1c). Conversely, K+ channel openers activate KATP channels and inhibit insulin secretion irrespective of glucose metabolism. Mutations that in result in reduced KATP channel activity, cause a beta-cell depolarization that is independent of the plasma glucose concentration, and give rise to congenital hyperinsulinism of infancy in humans (Fig. 1d). Conversely, mutations that impair beta-cell metabolism, or the ability of the KATP channel to respond to metabolically generated changes in nucleotide concentrations, decrease insulin secretion and lead to neonatal diabetes and maturity-onset diabetes of the young (Fig. 1e). They may also predispose to type 2 diabetes. In addition to beta cells, KATP channels are found in many other cell types, in which they play a multitude of physiological roles. These have been reviewed recently elsewhere [3]. This chapter focuses principally on our current knowledge of the beta-cell KATP channel in health and disease. Where appropriate, it also discusses the extra-pancreatic effects of disease-causing mutations in the genes encoding beta-cell KATP channels.

Molecular Structure The KATP channel consists of two types of protein subunit, Kir6.2 and SUR, which coassemble in a 4 : 4 stoichiometry to form an octameric complex (Fig. 2) [4]. Neither subunit can reach the plasma membrane in the absence of its partner, because each possesses an endoplasmic reticulum retention motif that must be masked by the other subunit [5]. This ensures that only fully functional KATP channels are trafﬁcked to the surface membrane. The K+-selective pore of the KATP channel is built up of four Kir6.2 (usually) or Kir6.1 (rarely) subunits (Fig. 2) [3]. Kir6.2 (KCNJ11) is a member of the inward rectiﬁer channel family [6,7] and possesses two transmembrane domains and cytosolic amino and carboxy termini. The site to which ATP binds to inhibit the channel lies on Kir6.2 [8]. Thus, there are four binding sites for ATP per KATP channel, each of which lies at the interface between the cytosolic domains of adjacent subunits (Fig. 2). However, binding of a single ATP is sufﬁcient to close the channel [9]. SUR, the sulfonylurea receptor, belongs to the ATP-binding cassette (ABC) protein superfamily [10,11]. It has 17 transmembrane helices arranged in three domains (TMD0, TMD1, and TMD2) which comprise ﬁve, six, and six helices, respectively. The two large cytosolic loops which follow TMD1 and TMD2 contain the nucleotide-binding domains NBD1 and NBD2,

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Fig. 2. a Representation of the transmembrane topology of a single Kir6.x (left) or SURx (right) subunit. Mg-nucleotide binding/hydrolysis at the nucleotide-binding domains (NBD1, NBD2) of SUR stimulates channel activity. Binding of adenosine triphosphate (ATP) or adenosine diphosphate (ADP) to Kir6.2 closes the pore, an effect that does not require Mg2+. b Images of the puriﬁed KATP channel complex obtained by single particle analysis of electron microscope images [14]. The colors indicate regions of different density (blue, strongest density). The images illustrate two slices through the structure towards the internal and external ends of the channel complex. c Top view of the EM density with models of Kir6.2 (blue) and SUR1 inserted [14]. ATP molecules are shown in green

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respectively (Fig. 2). These appear to associate in a sandwich dimer conformation, in which residues from the Walker A and B motifs of one NBD interact with the signature sequence of the other NBD. They thereby form two separate sites, with distinct properties, that bind and hydrolyze MgATP. MgATP hydrolysis has been demonstrated both for the isolated NBDs [12,13] and for the puriﬁed KATP channel complex [14]. In electrophysiological studies, both MgATP and MgADP can stimulate KATP channel activity [15–17]. However, evidence suggests that it is the presence of MgADP at site 2 (NBD2) that results in KATP channel opening [13,18], and that MgATP must be hydrolyzed to MgADP to produce channel activation [13]. SUR thus functions as a second metabolic sensor (in addition to Kir6.2), endowing the KATP channel with an exquisite sensitivity to changes in adenine nucleotide concentrations. It also serves as a target for both the inhibitory sulfonylurea drugs and the stimulatory K-channel openers, which act by binding to SUR, and thereby inﬂuence opening and closing (gating) of the Kir6.2 pore [19]. There are two genes that encode SUR, ABCC8 (SUR1) and ABCC9 (SUR2) [10,20,21]. The latter exists in several splice variants, the most important being SUR2A and SUR2B. SUR1 is expressed in pancreas and brain, SUR2A in cardiac and skeletal muscle, and SUR2B in a variety of tissues including brain and smooth muscle [22]. Differences in their SUR subunits contribute to the variable sensitivities of KATP channels in different tissues to cell metabolism and drugs.

Metabolic Regulation of KATP Channel Activity The regulation of the KATP channel is extremely complex for it is the target of numerous cytosolic chemicals and proteins. However, adenine nucleotides appear to be largely responsible for mediating the effects of glucose metabolism. Channel activity is determined by the balance between ATP inhibition (via Kir6.2) and MgATP/MgADP activation (via SUR1) [8]. The former predominates at low glucose concentrations, where metabolism is low, whereas the latter is favored when glucose metabolism is stimulated. Recent studies on the ATP sensitivity of KATP channels in intact and permeabilized beta cells support the view that the principal metabolic regulators are adenine nucleotides [23]. One of the classical arguments against ATP as the principal regulator of channel activity was that in inside-out membrane patches, half-maximal inhibition by ATP occurred at ATP concentrations (10 µM) far lower than those found in beta cells, and that the channel was largely blocked at ATP concentrations normally found in beta cells (1–10 mM) [24]. This conundrum has

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been resolved by the recognition that MgADP enhances channel activity, shifting it into the physiological range of ATP concentrations [23,25,26], and that even in the absence of glucose the KATP channel is rarely open so that glucose-dependent changes in KATP conductance involve only 5%–25% of the total current [2,23]. Many other compounds have been demonstrated to modulate the ATP sensitivity of the KATP channels, including phosphatidylinositol phosphates (PIP2, PIP3) and long-chain acyl-CoA esters [27–30]. While PIP2 and PIP3 are unlikely to be involved in glucose-dependent regulation of the KATP channel [23], they may be involved in mediating the effects of hormones and transmitters [28].

Regulation of KATP Channel Activity by Sulfonylurea Drugs Sulfonylureas have been used to treat type 2 diabetes for more than 50 years. Binding of sulfonylureas to SUR1 results in KATP channel closure, and in beta cells leads to membrane depolarization and insulin secretion (Fig. 1c) [19]. It has been proposed that sulfonylureas also have a direct effect on the betacell exocytotic apparatus [31], although this is contested [32]. Sulfonylureas can be divided into two distinct groups based on their afﬁnity for SUR2 [19,33]. One set of compounds, which includes glibenclamide and glimepiride, show high afﬁnity for both SUR1 and SUR2. Another group, which includes tolbutamide, gliclazide, and nateglinide, exhibits high afﬁnity only for SUR1. Intuitively, one might expect that the latter drugs would be preferable clinically as they would be expected to show greater beta-cell speciﬁcity. However, the UKPDS found no difference in mortality or morbidity between patients randomly assigned to glibenclamide or chlorpropamide (which structurally resembles tolbutamide) [34]. In inside-out membrane patches, high-afﬁnity block of KATP currents by sulfonylureas is not complete and 20%–40% of the current remains unblocked [35]. Intracellular MgADP increases the apparent efﬁcacy of sulfonylureas on Kir6.2/SURI channels, which explains why sulfonylureas block beta-cell KATP channels completely in intact cells [35]. The molecular mechanism of this effect remains unresolved. One possibility is that sulfonylureas displace MgADP from the NBDs of SUR and thus unmask the inhibitory effect of the nucleotide that is mediated via Kir6.2 subunit. Alternatively, MgADP may affect the mechanism by which sulfonylurea binding is translated into channel closure. The glinides (e.g., repaglinide) are derived from the benzamido moiety of glibenclamide, so it is perhaps not surprising that they also exert their thera-

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peutic effects via interaction with SUR [36]. Like the sulfonylureas, they can be divided into two groups based on their selectivity for SUR; however, the binding sites for sulfonylureas and glinides are not identical [19].

KATP Channels and Insulin Secretory Disorders In the absence of glucose, the resting potential of the pancreatic beta-cell is determined almost entirely by the activity of the KATP channel [2]. However, the open probability of the individual channel is very low, and it is estimated that only 5%–25% of the total KATP conductance is activated at rest [2]. There is also a highly non-linear relationship between the KATP channel conductance and the beta-cell membrane potential [23] (Fig. 3), which means that, close to the threshold for electrical activity, very small changes in KATP conductance have marked effects on electrical activity and insulin secretion. It can therefore be expected that tiny changes in KATP conductance resulting from mutations in KATP channel genes, or from impaired metabolic regulation of KATP channel activity, will cause diabetes or its converse, congenital hypoglycemia (Fig. 1) [68]. The study of monogenic forms of diabetes, and hyperinsulinemia, has conﬁrmed this idea and provides insight into the etiology of the more common, polygenic type 2 diabetes [22].

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Fig. 3. Relationship between membrane potential and KATP conductance. The gray shading indicates KATP conductance levels associated with electrical activity. After Tarasov et al. [23]

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Neonatal Diabetes Neonatal diabetes (ND) is deﬁned as diabetes presenting within the ﬁrst six months of life [22,37,38]. It affects about 1 in 200 000 live births. It is characterized by severe hyperglycemia, which may be either permanent (PNDM) or transient (TNDM), and in some cases is associated with mental and motor developmental delay, epilepsy, and muscle weakness (a condition known as DEND syndrome). Patients with intermediate DEND (iDEND) syndrome have neonatal diabetes coupled with delayed speech and walking, and may also show muscle weakness. Why some patients show transient neonatal diabetes, which may subsequently relapse, is not understood [39]. Likewise, it is unclear why carriers of some mutations do not develop overt diabetes (although they may have impaired glucose tolerance) or develop diabetes only later in life [40]. Obvious possibilities include the severity of the mutation, genetic background, and factors that enhance the risk of diabetes such as aging and obesity. Further clinical details are given in the chapter by Edghill and Hattersley).

Genetics of Neonatal Diabetes Neonatal diabetes is caused by mutations in a number of different genes, but most commonly in KCNJ11 (Kir6.2) and ABCC8 (SUR1) (chapter by Edghill and Hattersley)). Patients with ND-Kir6.2 mutations are all heterozygotes, whereas ND-SUR1 mutations are genetically more heterogeneous. At least for the Kir6.2 mutations, there is a reasonable correlation between genotype and the clinical phenotype, but there are several exceptions. To date, over 20 gain-of-function mutations in Kir6.2 associated with neonatal diabetes have been identiﬁed, the most prevalent being at residues R201 and V59 [38,41]. These mutations cluster around the putative ATP-binding site, or lie in regions of the protein involved in channel gating such as the slide helix, the cytosolic mouth of the channel, and the gating loops that link the ATP-binding site to the slide helix (Fig. 4). They may also affect residues involved in intrasubunit interactions. Functional analysis of ND-Kir6.2 mutations has provided useful insights into the working of the KATP channel. More recently, gain-offunction mutations have been also identiﬁed in the SUR1 subunit [42,43]. These are dispersed throughout the protein sequence but the ﬁeld is still too young to determine whether they cluster in any particular domains.

Functional Effects of Kir6.2 Mutations Causing Neonatal Diabetes The functional effects of more than 20 ND-Kir6.2 mutations have been explored by heterologous expression of recombinant channels [22,37–39,

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Fig. 4. a Structural model of Kir6.2 [57] viewed from the side. For clarity, only two transmembrane domains, and two separate cytosolic domains, are shown. Residues mutated in neonatal diabetes are shown in red. Adenosine triphosphate (ATP; green) is docked into its binding site. Of the residues implicated in neonatal diabetes R50, R201, Y330C, and F333I lie close to the ATP-binding site; F35, C42 and E332K at the interface between Kir6.2 subunits; G53 in a region postulated to interface with SUR1; and Q52, V59, C166, and I296L within regions of the channel involved in gating. b Structural model of Kir6.2 [57] viewed from above. For clarity, the transmembrane domains have been removed. Each subunit is colored a different color and ATP is shown in yellow, docked into its four ATP-binding sites. These lie at the interface between subunits. c Close-up of the putative ATP-binding site with residues lying within 3.5 Å of ATP indicated [57]. Residues mutated in neonatal diabetes are shown in red

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44–51]. Because all patients are heterozygotes, the heterozygous state has been simulated by coexpression of wild-type and mutant Kir6.2 with SUR1. Most studies have used Xenopus oocytes as the expression system of choice, because it is possible to ensure that the amount of wild-type and mutant mRNAs (but not protein) is identical. One caveat with these studies, therefore, is that channel assembly and/or the membrane trafﬁcking may differ in oocytes from that in pancreatic beta cells. However, recent studies in which wild-type and mutant (Kir6.2-R201H) channels were studied by stable transfection of INS-1 beta cells gave results similar to those found in oocytes [52]. All ND-Kir6.2 mutations analyzed to date produce an increase in the resting whole-cell KATP current and therefore constitute gain-of-function mutations [22,38]. The larger KATP current is predicted to reduce glucose-dependent depolarization of the beta-cell membrane, thus preventing activation of Ca2+ currents, electrical activity, Ca2+ inﬂux, and insulin secretion. In INS-1 beta cells expressing both wild-type and Kir6.2-R201H subunits (with SUR1) the membrane potential in glucose-free solution is unaltered [52], as expected since it is already dominated by the KATP conductance. However, metabolic substrates failed to depolarize the cell as completely, and action potential activity and insulin secretion were completely prevented [52]. The sulfonylurea tolbutamide, which closes KATP channels directly, restored both electrical activity and insulin release conﬁrming that it is the enhanced KATP current that is the problem. The increase in KATP current caused by ND-Kir6.2 mutations results from an impaired ability of MgATP to close the KATP channel. ATP concentration– response curves reveal that Kir6.2 mutations may shift the ATP concentration at which channel inhibition is half maximal to higher ATP concentrations and/or produce a “pedestal” of current that remains unblocked even at saturating ATP concentrations. In all cases, however, the magnitude of the KATP current at physiologically relevant ATP concentrations is increased. In beta cells, intracellular [ATP] ([ATP]i) in the presence of stimulatory glucose concentrations (i.e., >7 mM) ranges from around 2 mM to 10 mM [53]. Figure 5 shows that all ND-Kir6.2 mutations increase the KATP current at 3 mM MgATP at least 20-fold, and that those mutations which cause DEND syndrome have the greatest effect. There appears to be no correlation, however, between the magnitude of the KATP current and whether the mutation causes transient or permanent neonatal diabetes (Fig. 5).

Molecular Mechanism of Action of ND-Kir6.2 Mutations Analysis of gain-of function mutations in Kir6.2 has revealed they act by a variety of molecular mechanisms. First, all mutations impair the ability

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Fig. 5. a Fractional current remaining in the presence of 3 mM MgATP, measured in inside-out patches for wild-type (WT) KATP channels and KATP channels containing the indicated Kir6.2 mutations. Pale gray bars indicate mutations associated with transient neonatal diabetes (TNDM), dark gray bars mutations causing permanent neonatal diabetes (PNDM), hatched bars mutations causing intermediate DEND, and black bars mutations producing DEND syndrome (developmental delay, epilepsy, and neonatal diabetic syndrome). While it is likely that the developmental delay associated with the R50P mutation is due to the mutation (DEND syndrome), this cannot be conclusive as the patient had an episode of cerebral edema. The number of patches varied from 5 to 12. Data are taken from references [37,39,44–51]. b An estimate of the fractional whole-cell resting current remaining in the presence of 0.5 mM tolbutamide for wild-type (WT) KATP channels and KATP channels containing the indicated Kir6.2 mutations. Data were estimated by expressing the current in the presence of 3 mM azide and tolbutamide as a fraction of the current in azide alone. Pale gray bars indicate mutations associated with TNDM, dark gray bars mutations causing PNDM, and black bars mutations producing DEND syndrome. Asterisk, all patients with this mutation successfully transferred to sulfonylurea therapy; plus-sign, some patients transferred; open square, no patients transferred. The number of experiments varied from 5 to 17. Data are taken from references [37,39,44–51]

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of ATP to block the channel. This is evidenced by a reduced inhibitory effect of ATP in the absence of Mg2+ (ATP interacts only with Kir6.2, and not with SUR, in Mg-free solutions). Secondly, all mutations studied show increased activation by Mg-nucleotides. This manifests as a much greater decrease in channel ATP sensitivity on addition of Mg2+ than is observed for wild-type channels [46]. In the case of the R201H mutation, which has been most fully studied, activation by both nucleoside diphosphates and nucleoside triphosphates was markedly enhanced [50]. This indicates that NDKir6.2 mutations can also inﬂuence the functional coupling of SUR1 to Kir6.2. Neonatal diabetes mutations that lie within the putative ATP-binding site of Kir6.2 (e.g., R201, R50; Fig. 4) are considered to impair channel inhibition at Kir6.2 by interfering with nucleotide binding. While the electrophysiological data are entirely consistent with this view [37,44,48], this has yet to be demonstrated in biochemical studies. Many mutations, however, appear to affect ATP inhibition indirectly, by altering channel gating [44,45,47,51]. It is well established that mutations which increase the intrinsic channel open probability (i.e., that in the absence of ATP), reduce ATP inhibition [54,55]. The increase in intrinsic open probability is caused by stabilization of the open state of the channel, which shifts the gating equilibrium in the presence of ATP towards channel opening and thus indirectly reduces the channel ATP sensitivity. Many Kir6.2 mutations that cause neonatal diabetes affect the channel gating, shifting the channel towards the open state [44,45,47,51]. These mutations are found in regions of Kir6.2 considered to be involved in opening and closing of the pore. Thus, they lie at the cytosolic end of TM2, where the helices interact, which is postulated to form the inner gate of the channel (at least in the related bacterial channel KirBac1.1; [56]). They are also found in the slide helix, which is believed to slide along the plane of the membrane when the pore opens, and in the gating loops which connect the cytosolic domains to the slide helix [57]. Some mutations are also found at the interface between subunits, and molecular modeling suggests that these also move during channel gating [58]. No correlation between the molecular mechanism of action of Kir6.2 mutations and the severity of the clinical phenotype has been found. Indeed, both PNDM and DEND syndrome can result from mutations that impair channel gating [44,45,47,51] and this may also be true for mutations that alter ATP binding [48]. Rather, the data argue that it is not the type of the molecular mechanism but the magnitude of the KATP current at ATP concentrations found in the beta cell that determines the disease phenotype. The larger the current, the more severe the disease (Fig. 5).

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Functional Effects of SUR1 Mutations Causing Neonatal Diabetes To date, the functional effects of only a handful of SUR1 mutations associated with ND have been reported [42,43]. However, functional studies already indicate that ND-SUR1 mutations increase the KATP current in intact cells by a variety of molecular mechanisms. The ﬁrst to be described, F132L, resides within TMD0 in the cytosolic loop linking TMs 2 and 3 [42]. As TMD0 is known to modify the gating behavior of Kir6.2 [59], it is not perhaps surprising that the F132L mutation acts by stabilizing the open state of the KATP channel, and thereby indirectly impairing ATP inhibition. The other two NDSUR1 mutations that have been analyzed, I1424V and H1023Y, do not appear to alter the inhibitory effect of ATP, as evidenced by the fact that they have no effect on KATP channel ATP sensitivity in the absence of Mg2+ [43]. Nor do they alter the channel open probability. Instead, these mutations enhance the stimulatory effect of Mg-nucleotides. Residue I1424 lies within NBD2 and thus conceivably its mutation might affect ATP binding/hydrolysis. The molecular mechanism underlying the effect of H1023Y mutation is more difﬁcult to explain, as this residue lies in the extracellular linker between TMs 12 and 13. Further analysis of this mutation may therefore provide new insights into how SUR1 talks to Kir6.2.

Extra-Pancreatic Effects of ND Mutations Kir6.2 is widely expressed throughout the body, being found in heart, skeletal muscle, various endocrine cells, peripheral axons, and multiple types of brain neurons [6,7,60]. Gain-of-function mutations in Kir6.2 are therefore expected to reduce the electrical excitability of these cells also. This can explain the neurological and motor defects found in DEND patients. That much larger KATP currents appear to be required to affect extra-pancreatic tissues may reﬂect both differences in cell metabolism and in the complement of ion channels that regulate the cell membrane potential. The beta-cell is unusual in that [ATP]i ﬂuctuates in response to changes in external glucose, that KATP channels dominate the resting membrane potential, and that KATP channel activity is clearly sensitive to changes in extracellular glucose (in many cell types, KATP channels open only during ischemia). It is intriguing that although Kir6.2 is expressed in the heart [6,7] and is believed to contribute to the cardiac KATP channel [12,61], no obvious cardiac abnormalities have been reported in patients carrying Kir6.2 mutations. Similarly, mice engineered to carry gain-of-function Kir6.2 mutations that are expressed only in the heart are not substantially affected [62]. This conundrum may be resolved by the discovery that coexpression of Kir6.2 with the

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cardiac type of SUR (SUR2A), rather than SUR1, produces a very much smaller increase in the KATP current at physiologically relevant ATP concentrations [50]. This is because Mg-nucleotide activation is enhanced by SUR1 but not SUR2A [50,63]. It is unlikely that this is due to differences in the way that SUR2A couples to Kir6.2 because Mg-nucleotide activation is increased when Kir6.2 mutations are coexpressed with SUR2B (which differs from SUR2A only in its last 42 amino acids). We thus favor the idea that the differences in MgATP sensitivity between SUR1- and SUR2A-containing channels reﬂects the fact that SUR2A shows impaired MgATP hydrolysis [12]. Because SUR2A comprises the sulfonylurea receptor subunit in skeletal muscle KATP channels, it seems likely that, as in the heart, they will be unaffected by Kir6.2 mutations. The muscle weakness observed in DEND and iDEND patients must therefore be attributed to increased Kir6.2/SUR1 currents in the presynaptic nerve terminal, which reduce neurotransmitter release. The fact that a patient with an SUR1 mutation also exhibited muscle weakness [45] supports this hypothesis.

Implications for Therapy Before the discovery that ND can be caused by mutations in the KATP channel, it was assumed these patients suffered from type 1 diabetes with an unusually early onset, and they were treated with insulin. Several recent studies indicate that patients with Kir6.2 mutations that cause PNDM can be managed by sulfonylureas alone [64]. Importantly, not only is their quality of life enhanced but there is also a signiﬁcant improvement in their glucose control, with a marked decrease in HbA1C levels [64]. Figure 5b shows that tolbutamide, at a concentration that saturates the high-afﬁnity sulfonylurea-binding site on SUR1, reduces the whole-cell KATP current through channels carrying PNDM, TNDM, or iDEND mutations to a level that is similar to, or below, that seen for the wild-type channel in the absence of the drug. This explains why sulfonylureas are so effective in treating patients with these mutation. In contrast, sulfonylureas were not effective in patients with Kir6.2 mutations that cause DEND syndrome, presumably because these particular mutations strongly impaired the ability of sulfonylureas to block the KATP channel ([64]; Fig. 5b). A number of SUR1 mutations did not alter the sulfonylurea sensitivity of the recombinant channels, and patients carrying these mutations responded to sulfonylureas [43]. Clearly, insulin cannot ameliorate the extra-pancreatic effects of ND mutations, whereas sulfonylureas can close KATP channels in all tissues to which they have access. This raises the question of whether sulfonylureas may be beneﬁcial in treating the muscle weakness and developmental delay in iDEND patients, who carry sulfonylurea-sensitive Kir6.2 mutations. Given that

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cardiac KATP channels are barely affected by Kir6.2 mutations, and unaffected by SUR1 mutations, the question also arises as to whether it may better to treat ND patients with beta-cell speciﬁc sulfonylureas. This is because cardiac KATP channels are though to be important in the response to cardiac ischemia [3,61].

Congenital Hyperinsulinism of Infancy Congenital hyperinsulinism of infancy (CHI) is characterized by continuous, unregulated insulin secretion accompanied by severe hypoglycemia. The disease affects about 1 in 50,000 live births and presents at birth or within the ﬁrst year of life. Most CHI mutations are spontaneous (although familiar forms have also described) and homozygous, heterozygous, and compound heterozygous forms have been reported (reviewed in [41,65,66]). Congenital hyperinsulinism of infancy is often associated with neurological problems but it is not clear however whether these are caused by mutations themselves or whether they are a consequence of hypoglycemia.

Functional Effects KCNJ11 and ABCC8 are among the ﬁve different genes known to cause CHI, and about 50% of patients have mutations in ABCC8 (SUR1) [65]. More than 100 mutations have been described and these are distributed throughout the whole protein. Functionally, they may be divided into two classes. Class I mutations result in the loss of KATP channels in the plasma membrane, due to abnormalities in gene expression, protein synthesis, maturation, assembly or membrane trafﬁcking. Because SUR1 is required for correct trafﬁcking of Kir6.2 [5], both subunits fail to reach the membrane and are degraded. Class II mutations do not affect membrane targeting but impair the ability of the channel to respond to metabolic activators such as MgADP. Consequently the channels are always closed. In general, class I mutations cause a more severe phenotype while class II mutations result in a milder form of the disease. However, there is no precise genotype-phenotype correlation and the same mutation can cause CHI with different degrees of severity in different people. To date, fewer than 10 mutations causing CHI have been identiﬁed in Kir6.2 [22,41]. These decrease the magnitude of KATP current by reducing the channel density in the plasma membrane.

Type 2 Diabetes Islets isolated from type 2 diabetic patients show impaired insulin secretion in response to glucose [67]. However, in many patients sulfonylureas are still

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able to evoke insulin release. This raises the interesting possibility that type 2 diabetes may be associated with enhanced KATP channel activity, either because the channel fails to respond to metabolic signals, such as ATP, or because metabolism does not generate adequate ATP [68]. Indeed, the fact that mutations in KATP channel genes, and in metabolic genes, cause monogenic diabetes suggests that common polymorphisms in these genes may predispose to polygenic disease. Evidence in support of this idea is reviewed elsewhere [68]. A characteristic of type 2 diabetes is that it classically manifests in later life. This age dependence is widely believed to result from a reduction in mitochondrial function, as mitochondrial metabolism is known to decline with age and most of the ATP required for stimulus-secretion coupling in beta cells is mitochondrially generated [69]. These considerations therefore suggest that in type 2 diabetes, as in its monogenic counterpart, neonatal diabetes, the KATP channel may play an important role. Acknowledgments. Work in the authors’ laboratory is supported by the Wellcome Trust, the Royal Society and the European Union (Integrated Project Eurodia: LSHM-CT-2006-518153; Network of Excellence BioSim: LSHB-CT2004-005137). FMA is a Royal Society Research Professor. We thank our colleagues for their comments on earlier drafts of this chapter.

References 1. Ashcroft FM, Harrison DE, Ashcroft SJH (1984) Glucose induces closure of single potassium channels in isolated rat pancreatic beta cells. Nature 312:446–448 2. Ashcroft FM, Rorsman P (1989) Electrophysiology of the pancreatic beta-cell. Prog Biophys Mol Biol 54:87–143 3. Seino S, Miki T (2003) Physiological and pathophysiological roles of ATP-sensitive K+ channels. Prog Biophys Mol Biol 81:133–176 4. Inagaki N, Gonoi T, Seino S (1997) Subunit stoichiometry of the pancreatic beta-cell ATP-sensitive K+ channel. FEBS Lett 409:232–236 5. Zerangue N, Schwappach B, Jan YN, Jan LY (1999) A new ER trafﬁcking signal regulates the subunit stoichiometry of plasma membrane KATP channels. Neuron 22:537–548 6. Inagaki N, Tsuura Y, Namba N, Masuda K, Gonoi T, Horie M, Seino Y, Mizuta M, Seino S (1995) Cloning and functional characterization of a novel ATP-sensitive potassium channel ubiquitously expressed in rat tissues, including pancreatic islets, pituitary, skeletal muscle, and heart. J Biol Chem 270:5691–5694 7. Sakura H, Ämmalä C, Smith PA, Gribble FM, Ashcroft FM (1995) Cloning and functional expression of the cDNA encoding a novel ATP-sensitive potassium channel subunit expressed in pancreatic beta cells, brain, heart and skeletal muscle. FEBS Lett 377:338–344 8. Tucker SJ, Gribble FM, Zhao C, Trapp S, Ashcroft FM (1997) Truncation of Kir6.2 produces ATP-sensitive K channels in the absence of the sulfonylurea receptor. Nature 387:179–181

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9. Markworth E, Schwanstecher C, Schwanstecher M (2000) ATP4− mediates closure of pancreatic beta-cell ATP-sensitive potassium channels by interaction with 1 of 4 identical sites. Diabetes 49:1413–1418 10. Aguilar-Bryan L, Nichols CG, Wechsler SW, Clement JP IV, Boyd AE III, Gonzalez Herrerasosa H, Nguy K, Bryan J, Nelson DA (1995) Cloning of the beta-cell high-afﬁnity sulphonylurea receptor: a regulator of insulin secretion. Science 268: 423–425 11. Aguilar-Bryan L, Bryan J (1999) Molecular biology of adenosine triphosphate sensitive potassium channels. Endocr Rev 20:101–135 12. Masia R, Enkvetchakul D, Nichols CG (2005) Differential nucleotide regulation of KATP channels by SUR1 and SUR2A. J Mol Cell Cardiol 39:491–501 13. Zingman LV, Alekseev AE, Bienengraeber M, Hodgson D, Karger AB, Dzeja PP, Terzic A (2001) Signaling in channel/enzyme multimers: ATPase transitions in SUR module gate ATP-sensitive K+ conductance. Neuron 31:233–245 14. Mikhailov MV, Campbell JD, de Wet H, Shimomura K, Zadek B, Collins RF, Sansom MSP, Ford RC, Ashcroft FM (2005) 3-D structural and functional characterization of the puriﬁed KATP channel complex Kir6.2-SUR1. EMBO J 24:4166–4175 15. Nichols CG, Shyng SL, Nestorowicz A (1996) Adenosine diphosphate as an intracellular regulator of insulin secretion. Science: 272:1785–1787 16. Gribble FM, Tucker SJ, Ashcroft FM (1997) The essential role of the Walker A motifs of SUR1 in KATP channel activation by Mg-ADP and diazoxide. EMBO J 16: 1145–1152 17. Gribble FM, Tucker SJ, Haug T, Ashcroft FM (1998) MgATP activates the beta cell KATP channel by interaction with its SUR1 subunit. Proc Natl Acad Sci USA 95:7185–7190 18. Reimann F, Gribble FM, Ashcroft FM (2000) Differential response of KATP channels containing SUR2A or SUR2B subunits to nucleotides and pinacidil. Mol Pharmacol 58:1318–1325 19. Gribble FM, Reimann F (2003) Sulphonylurea action revisited: the post-cloning era. Diabetologia 46:875–891 20. Inagaki N, Gonoi T, Clement JP, Wang CZ, Aguilar-Bryan L, Bryan J, Seino, S (1996) A family of sulfonylurea receptors determines the properties of ATP-sensitive K+ channels. Neuron 16:1011–1017 21. Isomoto S, Kondo C, Yamada M, Matsumoto S, Higashiguchi O, Horio Y, Matsuzawa Y, Kurachi Y (1996) A novel sulfonylurea receptor forms with BIR (Kir6.2) a smooth muscle type ATP-sensitive K+ channel. J Biol Chem 271:24321–24324 22. Ashcroft, FM (2005) ATP-sensitive potassium channelopathies: focus on insulin secretion. J Clin Invest 115:2047–2058 23. Tarasov AI, Girard CAJ, Ashcroft FM (2006) ATP sensitivity of the ATP-sensitive K+ channel in intact and permeabilized pancreatic beta cells. Diabetes 55:2446–2454 24. Niki I, Ashcroft FM, Ashcroft SJH (1989) The dependence on intracellular concentration of ATP-sensitive K-channels and of Na-K-ATPase in intact HIT-T15 beta cells. FEBS Lett 257:361–364 25. Kakei M, Kelly RP, Ashcroft SJH, Ashcroft FM (1986) The ATP-sensitivity of K+ channels in rat pancreatic beta cells is modulated by ADP. FEBS Lett 208:63–66 26. Dunne MJ, Petersen OH (1986) Intracellular ADP activates ATP-sensitive K+ channels in an insulin-secreting cell line. FEBS Lett 208:59–62 27. Fan Z, Makielski JC (1997) Anionic phospholipids activate ATP-sensitive potassium channels J Biol Chem 272:5388–5395 28. Baukrowitz T, Fakler B (2000) KATP channels: Linker between phospholipid metabolism and excitability. Biochem Pharmacol 60:735–740

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29. Larsson O, Deeney JT, Bränström R, Berggren PO, Corkey BE (1996) Activation of the ATP-sensitive K+ channels by long chain acyl-CoA. J Biol Chem 271:10623– 10626 30. Gribble FM, Proks P, Corkey BE, Ashcroft (1998) Mechanism of cloned KATP channel activation by oleyol-CoA. J Biol Chem 273:26383–26387 31. Eliasson L, Renstrom E, Ammala C, Berggren PO, Bertorello AM, Bokvist K, Chibalin A, Deeney JT, Flatt PR, Gabel J, Gromada J, Larsson O, Lindstrom P, Rhodes CJ, Rorsman P (1996) PKC-dependent stimulation of exocytosis by sulfonylureas in pancreatic beta cells. Science 271:813–815 32. GarciaBarrado MJ, Jonas JC, Gilon P, Henquin JC (1996) Sulphonylureas do not increase insulin secretion by a mechanism other than a rise in cytoplasmic Ca2+ in pancreatic B-cells. Eur J Pharmacol 298:279–286 33. Gribble FM, Tucker SJ, Seino S, Ashcroft FM (1998) Tissue speciﬁcity of sulphonylureas: studies on cloned cardiac and beta cell KATP channels. Diabetes 47:1412–1418 34. United Kingdom Prospective Diabetes Study (UKPDS) 13 (1995) Relative efﬁcacy of randomly allocated diet, sulphonylurea, insulin, or metformin in patients with newly diagnosed non-insulin dependent diabetes followed for three years. BMJ 310:83–88 35. Gribble FM, Tucker SJ, Ashcroft FM (1997) The interaction of nucleotides with the tolbutamide block of KATP currents: a reinterpretation. J Physiol 504:35–45 36. Dabrowski M, Wahl P, Holmes WE, Ashcroft FM (2001) Effect of repaglinide on cloned beta cell, cardiac and smooth muscle types of ATP-sensitive potassium channel. Diabetologia 44:747–756 37. Gloyn AL, Pearson ER, Antcliff JF, Proks P, Bruining JG, Slingerland AS, Howard N, Srinivasan S, Silva JMCL, Molnes J, Edghill EL, Frayling TM, Temple IK, Deborah Mackay D, Shield JPH, Sumnik Z, van Rhijn A, Wales JKH, Clark P, Gorman S, Aisenberg J, Ellard S, Njølstad PR, Ashcroft FM, Hattersley AT (2004) Activating mutations in the gene encoding the ATP-sensitive potassium-channel subunit Kir6.2 and permanent neonatal diabetes. N Engl J Med 350:1838–1849 38. Hattersley AT, Ashcroft FM (2005) Activating mutations in Kir6.2 and neonatal diabetes: new clinical syndromes, new scientiﬁc insights, and new therapy. Diabetes 54:2503–2513 39. Gloyn AL, Reimann F, Girard C, Edghill EL, Proks P, Pearson ER, Temple IK, Mackay DJG, Shield JPH, Freedenberg D, Noyes K, Ellard S, Ashcroft FM, Gribble FM, Hattersley AT (2005) Relapsing diabetes can result from moderately activating mutations in KCNJ11. Hum Mol Genet 14:925–934 40. Yorifuji T, Nagashima K, Kurokawa K, Kawai M, Oishi M, Akazawa Y, Hosokawa M, Yamada Y, Inagaki N, Nakahata T (2005) The C42R mutation in the Kir6.2 (KCNJ11) gene as a cause of transient neonatal diabetes, childhood diabetes, or later-onset, apparently type 2 diabetes mellitus. J Clin Endocrinol Metab 90:3174–3178 41. Gloyn AL, Siddiqui J, Ellard S (2006) Mutations in the genes encoding the pancreatic beta-cell KATP channel subunits Kir6.2 (KCNJ11) and SUR1 (ABCC8) in diabetes mellitus and hyperinsulinism. Hum Mutat 27:220–231 42. Proks P, Arnold AL, Bruining J, Girard C, Flanagan SE, Larkin B, Colclough K, Hattersley AT, Ashcroft FM, Ellard S (2006) A heterozygous activating mutation in the sulphonylurea receptor SUR1 (ABCC8) causes neonatal diabetes. Hum Mol Genet 15:1793–1800 43. Babenko AP, Polak M, Cavé H, Busiah K, Czernichow P, Scharfmann R, Bryan J, Aguilar-Bryan L, Vaxillaire M, Froguel P (2006) Activating mutations in the ABCC8 gene in neonatal diabetes mellitus. N Engl J Med 355:456–466

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44. Proks P, Antcliff JF, Lippiat J, Gloyn A, Hattersley AT, Ashcroft FM (2004) Molecular basis of Kir6.2 mutations associated with neonatal diabetes or neonatal diabetes plus neurological features. Proc Natl Acad Sci USA 101:17539–17544 45. Proks P, Girard C, Haider S, Gloyn AL, Hattersley AT, Sansom MSP, Ashcroft FM (2005) A novel gating mutation at the internal mouth of the Kir6.2 pore is associated with DEND syndrome. EMBO Rep 6:470–475 46. Proks P, Girard C, Ashcroft FM (2005) Functional effects of KCNJ11 mutations causing neonatal diabetes: enhanced activation by MgATP. Hum Mol Genet 14:2717–2726 47. Proks P, Girard C, Bævre H, Njolstad PR, Ashcroft FM (2006) Functional effects of mutations at F35 in the NH2-terminus of Kir6.2 (KCNJ11), causing neonatal diabetes, and response to sulphonylurea therapy. Diabetes 55:1731–1737 48. Shimomura K, Girard C, Proks P, Nazim J, Lippiat JD, Cerutti F, Lorini R, Ellard S, Hattersley AT, Barbetti F, Ashcroft FM (2006) Mutations at the same residue (R50) of Kir6.2 (KCNJ11) that cause neonatal diabetes produce different functional effects. Diabetes 55:1705–1712 49. Tammaro P, Girard C, Molnes J, Njølstad PR, Ashcroft FM (2005) Kir6.2 mutations causing neonatal diabetes provide new insights into Kir6.2-SUR1 interactions. EMBO J 24:2318–2330 50. Tammaro P, Proks P, Ashcroft FM (2006) Functional effects of naturally occurring KCNJ11 mutations causing neonatal diabetes on cloned cardiac KATP channels. J Physiol 571:3–14 51. Girard C, Shimomura K, Proks P, Absalom N, de Nanclares PG, Ashcroft FM (2006) Functional analysis of six Kir6.2 (KCNJ11) mutations causing neonatal diabetes. Pﬂugers Arch 453:323–332 52. Tarasov A, Welters HJ, Senkel S, Ryffel GU, Hattersley AT, Morgan NG, Ashcroft FM (2006) A Kir6.2 mutation causing neonatal diabetes impairs electrical activity and insulin secretion from INS-1 beta cells. Diabetes 55:3075–3082 53. Erecinska M, Bryla J, Michalik M, Meglasson MD, Nelson D (1992) Energy-metabolism in islets of Langerhans. Biochim Biophys Acta 1101:273–295 54. Trapp S, Proks P, Tucker SJ, Ashcroft FM (1998) Molecular Analysis of KATP channel gating and implications for channel inhibition by ATP. J Gen Physiol 112:333–349 55. Enkvetchakul D, Loussouarn G, Makhina E, Nichols CG (2001) ATP interaction with the open state of the KATP channel. Biophys J 80:719–728 56. Kuo AL, Gulbis JM, Antcliff JF, Rahman T, Lowe ED, Zimmer J, Cuthbertson J, Ashcroft FM, Ezaki T, Doyle DA (2003) Crystal structure of the potassium channel KirBac1.1 in the closed state. Science 300:1922–1926 57. Antcliff JF, Haider S, Proks P, Sansom MSP, Ashcroft FM (2005) Functional analysis of a structural model of the ATP-binding site of the KATP channel Kir6.2 subunit EMBO J 24:229–239 58. Haider S, Grottesi A, Hall BA, Ashcroft FM, Sansom MSP (2005) Conformational dynamics of the ligand-binding domain of inward rectiﬁer K channels as revealed by molecular dynamics simulations: Toward an understanding of Kir channel gating. Biophys J 88:3310–3320 59. Babenko AP, Bryan J (2003) SUR domains that associate with and gate KATP pores deﬁne a novel gatekeeper. J Biol Chem 278:41577–41580 60. Karschin C, Ecke C, Ashcroft FM, Karschin A (1997) Overlapping distribution of KATP channel-forming Kir6.2 subunit and the sulfonylurea receptor SUR1 in rodent brain. FEBS Lett 401:59–64 61. Flagg TP, Nichols CG (2005) Sarcolemmal KATP channels: what do we really know? J Mol Cell Cardiol 39:61–70

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62. Koster JC, Knopp A, Flagg TP, Markova KP, Sha Q, Enkvetchakul D, Betsuyaku T, Yamada KA, Nichols CG (2001) Tolerance for ATP-insensitive K-ATP channels in transgenic mice. Circ Res 89:1022–1029 63. Matsuoka T, Matsushita K, Katayama Y, Fujita A, Inageda K, Tanemoto M, Inanobe A, Yamashita S, Matsuzawa Y, Kurachi Y (2000) C-terminal tails of sulfonylurea receptors control ADP-induced activation and diazoxide modulation of ATP-sensitive K+ channels. Circ Res 87:873–880 64. Pearson ER, Flechtner I, Njolstad PR, Malecki MT, Flanagan SE, Larkin B, Ashcroft FM, Klimes I, Codner E, Iotova V, Slingerland AS, Shield J, Robert J, Holst JJ, Clark PM, Ellard S, Sovik O, Polak M, Hattersley AT (2006) Switching from insulin to oral sulfonylureas in patients with diabetes due to Kir6.2 mutations. N Engl J Med 355: 467–477 65. Dunne MJ, Cosgrove KE, Shepherd RM, Aynsley-Green A, Lindley KJ (2004) Hyperinsulinism in infancy: from basic science to clinical disease. Physiol Rev 84:239–275 66. Glaser B (2000) Hyperinsulinism of the newborn. Semin Perinatol 24:150–163 67. Del Guerra S, Lupi R, Marselli L, Masini M, Bugliani M, Sbrana S, Torri S, Pollera M, Boggi U, Mosca F, Del Prato S, Marchetti P (2005) Functional and molecular defects of pancreatic islets in human type 2 diabetes. Diabetes 54:727–735 68. Ashcroft FM, Rorsman P (2004) Molecular defects in insulin secretion in type-2 diabetes. Rev Endocr Metab Disord 5:135–142 69. Wollheim CB, Maechler P (2002) Beta-cell mitochondria and insulin secretion— Messenger role of nucleotides and metabolites. Diabetes 51 (Suppl 1):S37–S42

21. Glucokinase in Glucose Homeostasis, Diabetes Mellitus, Hypoglycemia, and as Drug Receptor Franz M. Matschinsky

Summary. The pivotal role of the glucose phosphorylating enzyme glucokinase in glucose homeostasis is discussed, including its established functions as glucose sensor in pancreatic beta cells and as control step in hepatic carbohydrate metabolism. A brief review of “Glucokinase Disease,” a group of hyper- and hypoglycemic clinical phenotypes caused by mutations of the glucokinase gene and inherited as an autosomal dominant trait, and a concise assessment of a newly discovered class of promising antidiabetic drugs that activate glucokinase allosterically, round out this perspective chapter.

Introduction The central role of the glucose phosphorylating enzyme glucokinase (GK) in glucose homeostasis, diabetes mellitus, and hypoglycemia has been discussed thoroughly in several recent reviews and two authoritative monographs [1– 5]. The discovery of GK activator drugs (GKAs) and their potential as new antidiabetic medication has also been the topic of up-to-date publications [6–9]. Several chapters in this volume deal with various issues of stimulus secretion coupling in pancreatic islet cells activated by fuels including glucose and do thus complement the discussion to follow. The present brief perspective was written with this background in mind, highlighting newer developments and ideas inﬂuencing the ﬁeld but with only limited reference to the older literature except where essential for reasons of clarity. Department of Biochemistry and Biophysics, University of Pennsylvania, School of Medicine, 501 Stemmler Hall, 36th & Hamilton Walk, Philadelphia, PA 19104, USA

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Outline of the Glucokinase Glucose Sensor Concept Glucose homeostasis in man with his physiological setpoint of about 5 mM is best comprehended in engineering terms by envisioning a glucostat system consisting of a glucose sensor device, an effective signaling capability, and mechanisms for rapid adjustment of blood sugar levels deviating from this setpoint [1,10,11]. Extrapolating from engineering to biology the following picture emerges. The pancreatic islets of Langerhans serve as a glucostat; they contain glucokinase which operates as the glucose sensor element; islet cells secrete insulin and glucagon as hormonal signals; and liver and muscle function as insulin and glucagon regulated sink or source of glucose. In the aggregate, the islets of Langerhans perform as one single endocrine master gland

Fig. 1. Glucose and glucokinase activator (GKA) actions on network of sentinel glucokinase (GK) cells and parenchymal hepatocytes in glucose homeostasis. Glucose alone or enhanced by GKAs act on GK in a wide variety of glucose sensor cells, which results in secondary signaling throughout the network via hormones or the autonomic nervous system impacting on glucose homeostasis. GKAs and glucose also affect the metabolism of hepatoparenchymal cells, resulting in enhanced clearance of blood glucose and glycogen synthesis. It should be appreciated that glucose or GKAs may result in stimulatory or inhibitory effects, directly or indirectly. For example they stimulate beta cells but inhibit alpha cells, directly or indirectly. For reasons of clarity plus/minus signs indicating such actions are omitted. Reproduced from Matschinsky et al. [1] with permission

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in this feedback system. All islet beta cells together amount to the overriding determinant of the glucose setpoint and constitute the backbone of glucose homeostasis in man. Considering the focus of this perspective, it is emphasized here that even minor activation or inactivation of GK by itself results in lowering or raising of the glucose setpoint. In reality, the situation is more complex as illustrated by the following example: additional GK containing cells exist forming a complex network of glucose sensor cells (Fig. 1). This includes enteroendocrine, hypothalamic, hepatic vascular, and anterior pituitary cells, which are thought to inﬂuence the basic regulatory system in many different ways [1,12,13]. It is remarkable that these GK-based glucose sensor cells contain probably less than 1% of the total complement of the bodies GK. The remaining 99% of the enzyme resides in the parenchymal cells of the liver and forms the basis for a high capacity process to clear glucose from the blood and store it in the form of glycogen. Hepatic parenchymal GK does therefore not operate as a glucose sensor element in the strict meaning of the word, and alterations of its activity have only a limited impact on the bodies glucose setpoint.

Prototypical Design Features of Pancreatic Beta Cells as Glucose Sensor Cells The glucose responsive, insulin-producing pancreatic beta cell is conceptualized as the archetypical glucose sensor cell [1]. It has certain essential design features: (1) it contains a store of large, dense core granules of the hormone insulin, i.e., it has the capability of generating a hormonal signal; (2) it is endowed with high capacity glucose transporters which facilitate the rapid equalization of intra- and extracellular glucose concentrations; (3) it contains GK which has kinetic characteristics allowing it to translate physiological glucose level changes into corresponding changes of metabolic ﬂuxes and energy potentials; (4) it exhibits constitutive GK expression which is glucose inducible in a manner largely independent of insulin; and (5) it contains in its cell membrane potassium and calcium channels that are regulated by metabolic and membrane potential changes constituting the molecular basis for cellular excitability and membrane depolarization, which results in elevated intracellular free calcium levels and stimulation of hormone exocytosis. It would be highly misleading to extrapolate from this particular case of the pancreatic beta cell that other GK containing glucose sensor cells are similarly designed. For example [12,13], GK was recently discovered in the FSH (follicle stimulating hormone) and LH (luteinizing hormone) secreting gonadotropes of the anterior pituitary indicating glucose sensor capacity of these cells (Fig. 2). Hypothalamic gonadotropin releasing hormone (GnRH) induces pulsatile

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Fig. 2. Glucose sensor cells of the anterior pituitary. Higher magniﬁcation images from a rat pituitary double-labeled with antibodies to glucokinase (red) and either FSH (follicle stimulating hormone), LH (luteinizing hormone), or TSH (thyroid stimulating hormone) (green). The staining intensity of glucokinase in the large cells was strongly correlated with that for FSH (top). This was not true for LH where many large cells intensely stained for glucokinase were lightly stained for LH (see cells in the lower left corner, middle). Most of the thyrotropes have less intense glucokinase staining, although occasional cells have more intense staining (see cells in the upper right corner, bottom). Bar: 10 µm. Reproduced from Sorenson et al. [13] with permission

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hormone release from gonadotropes associated with oscillatory hyperpolarization rather than depolarization of the cell membrane as manifestation of a molecular process of stimulus secretion coupling that is entirely different from that of the pancreatic beta cell [14–16]. GnRH activates a Gq-coupled (G-protein q coupled) membrane receptor and stimulates phospholipase C (PLC) leading to the generation of the second messengers IP3 (inositol trisphosphate) and diacylglycerol (DAG), which then cause calcium release from internal stores and protein kinase C (PKC) activation, respectively, with the end result of increased FSH/LH secretion. It is hypothesized that glucose, mediated by GK, modiﬁes or may even be required to permit this stimulation process in analogy to the glucose requirement for GLP-1 and acetylcholine action on beta cells [13]. It is also attractive to speculate that metabolically generated DAG might be the mediator. Cellular DAG levels may be elevated by high glucose because fatty acid oxidation is usually blocked and lipogenesis augmented as a result of a rising Phosphate-potential (including a drop of 5′-AMP) and increased malonyl-CoA. We have recently proposed that this direct action of glucose at the level of the anterior pituitary could play a role in the in the development of nutritional infertility [13].

The Unique Biochemistry of Hepatic Glucokinase and Its Physiological Significance As stated earlier, 99% of the body’s GK resides in the hepatocytes. This is explained by the large liver mass and by the manifold higher concentration of GK in liver cells as compared to other GK containing cells. Even though not critical for establishing the 5 mM setpoint of the organism (the primary responsibility of the endocrine cells of the islets of Langerhans), hepatic GK plays a signiﬁcant role in the regulation of carbohydrate metabolism [17–19]. First, the concentration of hepatic GK is determined by insulin owing to the liver-speciﬁc expression control of the enzyme at the transcriptional level, as illustrated by the observation that diabetes or total starvation result in marked reduction of GK mRNA and protein in the liver [19]. Second, hepatic GK is subject to short-term control by GK regulatory protein (GKRP), in contrast to GK regulation in islet cells which lack GKRP [1,17,18]. Glucokinase regulatory protein, a 68 kDa nuclear protein, mediates the nuclear sequestration of GK when glucose levels are low. However, when glucose levels rise postprandially the nuclear GK/GKRP complex dissociates because glucose is a competitive antagonist of GKRP. Glucokinase is therefore released into the cytosol and glucose phosphorylation is stimulated, facilitating glycogen synthesis and curbing gluconeogenesis.

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Mutations in the Glucokinase Gene Result in Autosomal Dominantly Inherited “Glucokinase Disease” The GK glucose sensor paradigm implies that even small changes of the enzymatic capacity of GK in the beta cells would result in changes of the glucose threshold of glucose stimulated insulin release (GSIR) and alterations in glucose homeostasis, i.e., diabetes mellitus or hyperinsulinemic hypoglycemia. This prediction was conﬁrmed by the discoveries that inactivating mutations of GK do indeed cause diabetes mellitus of the MODY type (Maturity Onset Diabetes of the Young) or PNDM (Permanent Neonatal Diabetes Mellitus), and that activating mutations of the enzyme result in Persistent Hyperinsulinemic Hypoglycemia of Infancy (PHHI) [1–3,20–30]. An estimated 250 mutations of the GK gene have been identiﬁed in the last 15 years in patients with diabetes or hypoglycemia and it can be predicted that many more will be detected. A large selection of missense mutations among these has been studied using recombinant mutant enzyme in comparison to wild type. Most of these investigations were performed with GST-GK (Glutathionyl-S-Tranferase–GK) fusion protein. The strategy was validated by showing that the kinetic constants of GST-tagged enzymes are virtually indistinguishable from those of cleaved, pure GK in 21 of 22 cases (Y214A/V452A representing the one exception which exhibited an unexplained decrease of the Kcat by 80% when GST was removed). This characterization included determinations of kinetic constants of the enzyme [23–28,31], tests of thermal stability [23–28,31], assessment of the response of GK to GKRP and allosteric GK activator drugs [26,31], measurements of equilibrium glucose binding or of glucose binding kinetics using tryptophan ﬂuorescence [29,32,33], and cell biological evaluation of enzyme characteristics using viral vector technologies or gene targeting methods in mice [30,34]. A great wealth of information about the enzyme per se and its role as glucose sensor has been gathered by these efforts. In the vast majority of cases it was possible to use the results of these studies to explain the clinical phenotype resulting from the genetic lesions. Using minimal mathematical modeling, it was possible to compute the glucose thresholds for GSIR in GK-linked hypo- and hyperglycemia cases in reasonable agreement with the observed clinical and laboratory results [35]. Glucokinase gene mutations were found to change the patient’s glucose setpoints over a wide concentration range from 1 mM to as high as 50 mM or even higher, owing primarily to the effects on pancreatic beta cells. In the course of these studies it was discovered that the majority of activating mutations causing PHHI were clustered in a circumscribed domain of the enzyme located at a 20 Å´ distance away from the substrate binding site, suggesting the existence of a hitherto unknown allosteric activator region of GK [23–26]. Activating mutations increase the glucose afﬁnity of the enzyme, lower the

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Hill coefﬁcient with regard to glucose as substrate, may increase the Kcat, and may also prevent inhibition by GKRP and block activation by GKAs. They may increase the quantum yield (Q) of tryptophan ﬂuorescence, a measure of structural compactness of a protein molecule [29]. These new results provide spectroscopic evidence supporting the proposal derived from crystallographic [8] and kinetic studies [32,33,36] that the enzyme does exist at least in three conformations (termed the superopen, the open, and the closed forms) and that the ligand induced transition between these forms might explain the cooperativity of GK with regard to glucose. The lowest quantum yield is obtained in the absence of glucose and saturating levels of glucose increase Q about twofold. Certain activating mutants characterized by high glucose afﬁnity, a low Hill coefﬁcient and, not infrequently, by refractoriness to GKRP and GKAs (GK activators) show a relatively high Q in the absence of glucose, suggesting an intermediate conformational state, perhaps equivalent to the hypothetical open form. It is important to appreciate that some hyper- and hypoglycemic human phenotypes were not fully explained by the results of biochemical and biophysical analysis of the corresponding recombinant protein. It is possible that unidentiﬁed characteristics of glucokinase are signiﬁcantly modiﬁed by such mutations leading to a pathological phenotype [12]. If this were true, gene targeting studies in mice using the mutant human enzyme in question promise to provide new insights [30].

Discovery and Potential of GKAs for Treatment of Type 2 Diabetes Mellitus When the biochemical basis of PHHI linked to GK was elucidated, culminating in the recognition that activating mutations do cluster in a putative allosteric activator site, it was discovered that micromolar concentrations of propionic acid amide derivatives and of small molecules with other chemical core structures were able to activate glucokinase by binding highly speciﬁcally to the same domain of the enzyme and causing the same kinetic changes as the activating mutations [1,6–9,37]. This was accomplished because the availability of GKAs facilitated the crystallization of GK complexed with glucose and GKA allowing the identiﬁcation of the contact points of the activators (Fig. 3). It is noteworthy that GKAs are unable to bind to GK in the absence of glucose. This can be demonstrated by the absence of tryptophan ﬂuorescence enhancement when GKAs are added to an enzyme solution. The inability of GK to bind the activator is explained by the structural features of the superopen conformation of the enzyme in which the drug binding site is unraveled and/or access of the drug impeded. Glucose induces

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Fig. 3A–D. Structure of allosteric GKA binding site of glucokinase and examples of GKA effects on normal and mutant enzymes. Panel A conveys an impression of the structure of the allosteric activator site of GK and shows contact amino acids for GKAs. Panel B shows the concentration-dependent effect of a GKA on GK activity as a function of glucose in the assay. Note that the relative effect of the activator is highest at the lowest glucose concentrations. Panel C gives examples of mutants that are refractory to the action of GKAs whereas panel D illustrates the effectiveness of GKAs in reversing the inhibition resulting from inactivating mutations. Reproduced from Matschinsky et al. [1] with permission

a conformation which gives rise to the drug binding site. It is noteworthy mechanistically that the activating mutant W99R binds GKAs in the absence of glucose as indicated by an increase of tryptophan ﬂuorescence [29]. It has been speculated that liver and pancreatic islet tissue might contain a natural GK activator. The existence of such an endogenous GK activator would make much sense biologically and great efforts should be expended in searching for it. As one might predict from the physiological and genetic information discussed in the preceding paragraphs, GKAs do lower the threshold for GSIR of pancreatic islets as a consequence of increased glucose metabolism, an elevated phosphate-potential of beta cells, and a rise of free intracellular calcium in the presence of glucose [6,7,9]. Glucokinase activators stimulate

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glycogen synthesis and curb gluconeogenesis in the liver, as well as lowering blood sugar in normal and diabetic laboratory animals. They prevent the development of hyperglycemia in the course of diet induced obesity (DIO) without inﬂuencing weight gain. It can be anticipated that they will be similarly effective in diabetic humans provided some residual functional capacity of the beta cell is preserved. Since GKAs also improve hepatic carbohydrate metabolism provided hepatocytes contain a critical level of GK, additional beneﬁts can be predicted to arise from this second site of drug action. One should also consider the possibility that GKAs could inﬂuence the GK containing cells of the complex network of glucose sensors and impact on glucose homeostasis in ways to be elucidated (Fig. 1). To gain a broader perspective regarding the proposed use of GKAs as antidiabetic drugs, it is perhaps instructive to draw attention to current rationales and strategies for treating vascular hypertension. The primary molecular cause of hypertension remains unknown as is the cause of type 2 diabetes. However, each disease has a characteristic dangerous “symptom,” high blood pressure in one and high blood glucose in the other. Both hypertension and hyperglycemia are, per se, pathogenic factors if untreated. Antihypertensive drug therapy takes advantage of many dissimilar mechanisms that lower blood pressure without clear knowledge about the root cause of the disease. Yet the strategy has been remarkably successful even though it “merely treats the symptom.” The situation in diabetes therapy is comparable. Defective GK is almost certainly not a prevalent molecular cause of classical type 2 diabetes mellitus. However, GK is an ideal drug target because it has a very high impact factor in glucose homeostasis [1] such that its activation results in lower blood sugar even though the cause of hyperglycemia might have little to do with this enzyme. Glucokinase activator drugs promise therefore to gain a prominent place among currently available antidiabetic medicines.

Challenges for Future GK Research It has been about 40 years since the presence and physiological signiﬁcance of GK of the liver and soon thereafter of the islets of Langerhans were discovered [38–41]. The interest in, and the knowledge about this enzyme have increased nearly exponentially over time, with a dramatic surge in research reports in the last decade. This should not be surprising since the importance of GK for glucose homeostasis is perhaps comparable to that of hemoglobin for oxygen physiology. The exploration of both molecules has resulted in insights of fundamental biological signiﬁcance and discoveries of great practical relevance, and there is no end in sight for future research developments for either one of these. Several GK related issues deserve special attention in

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the years to come: (1) the functional and structural basis of cooperative kinetics with respect to glucose; (2) expression control of GK in extrahepatic tissue; (3) the role of GK in gonadotropes and other extra-pancreatic glucose sensor cells; (4) the search for endogenous GK activator(s); (5) comprehensive studies of the biochemical and biophysical genetics of glucokinase disease in man; and (6) biochemical pharmacology of GKAs and evaluation of potential drugs for the treatment of type 2 diabetes mellitus. Acknowledgments. These studies were supported by the NIH (NIDDK ROI22122) and a grant from the American Diabetes Association.

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39. Sols A, Salas M, Vinuela E (1964) Induced biosynthesis of liver glucokinase. Adv Enzyme Regul 2:177–188 40. Sharma C, Manjeshwar R, Weinhouse S (1964) Hormonal and dietary regulation of hepatic glucokinase. Adv Enzyme Regul 2:189–200 41. Matschinsky FM, Ellerman JE (1968) Metabolism of glucose in the islets of Langerhans. J Biol Chem 243:2730–2736

Subject Index

a ABCC8 435, 438, 445 ACh 162 action potential 91 activated by 38 activin A 275 activin, retinoic acid 274 acylation 60 acyl CoA 59 acyl-CoA 227 adenine nucleotide translocase 60 adenine nucleotides 55–56 adeno-associated virus 289 adenoviruses 289 adipocyte-derived cytokines 224–225, 228 adult 13 AKAP 141 Akt 227–228, 231–232, 239, 242–243 allosteric GK activator drugs 456 allosteric GKA binding site 458 alpha cells 6 AMPK 158 amplifying action 161 amyloid 382 amyloid deposition 387 amyloid stress 225–226 amyloidosis 382 anaplerosis 59, 62 anterior pituitary cells 453 antidiabetic medicines 459 apamin 98

apoptosis 75–78, 80–82, 84, 215–218, 221–228, 230, 232–235, 237–238, 240, 243, 323, 385 arachidonic acid 101 archetypical glucose sensor cell 453 architecture 5 ATGs 223 ATP 241 ATP release 119 ATP/ADP 53, 66 ATP/ADP ratio 54, 56–57, 67 ATP-sensitive potassium channel see KATP channel autocata-lytic 38 autophage 215, 217, 222–223, 225, 228–229, 231 autophagic cell death 215, 217, 222–223, 229 autophagy 223, 225, 228–229, 237–238

b 6-Bnz-cAMP 136–137 7B2 40 BAD 230–231 basal release of insulin 124 baseline beta-cell mass 224 beta-catenin 230–231 beta-cell 75–84, 120 beta cell neogenesis 219–221, 236 beta cell replication 216–221, 229, 235 beta-cell death 42, 215–217, 222–225, 229–230, 232, 234, 240

465

466

Subject Index

beta-cell death-measurement 223 beta-cell function 382 beta-cell growth 215–219, 221–225, 227, 229–237, 239, 241, 243 beta-cell mass 215–226, 228, 234–236, 238, 242, 322, 382 beta-cell neogenesis 219–221, 236 beta-cell precursor 219 beta-cell proliferation 267 beta-cell regeneration 266, 272 beta-cell size 220–221, 231, 247 beta-cell stress 224–225 beta-cell transdifferentiation 220–221 beta-cell turnover 249 beta cells 6, 14, 18, 451 BETA2/NeuroD1 21 biochemical pharmacology 460 biologic activity 34 biophysical genetics 460 biosynthesis of insulin 31 biphasic pattern 150 birth weight 224 BK 95 BK channels 95 blind drunk mutation 167 bone marrow (BM) 272, 296 Brefeldin A 36

c C2 element 17 Ca2+-activated K+ current 96 Ca2+-activated K+ channels 96 Ca2+-activated potassium channels 92 Ca2+ channels 93, 431–432 Ca2+ oscillations 120, 125 Ca2− 57, 125, 225–228, 232, 240, 243 calcineurin 163 calcineurin/NFATc 255 calcium 40 cAMP 133–146, 243 cAMP analog 134, 136–138, 143–144 cAMP compartment 141 cAMP compartmentation 133, 141, 145 cAMP-GEF 133–137, 141, 146 cAMP-GEFII 135–140, 143 canine 5 carbonic anhydrase-II 219, 236 carboxypeptidase 37

carboxypeptidase B 37 carboxypeptidase E 41 caspase degenerative disease 222–223 cats 389 CaV1.1, 1.2, 1.3, and 1.4 channels 159 Cav1.2 93 Cav1.3 93 CaV2.1 (P/Qtype), 2.2 (N-type) and 2.3 (R-type) 159 Cdx2/3 18 cell aggregation 112 cell communication 112 cell composition 9 cell-attached capacitance measurements 164 cell-replacement therapy 304 ceramide 390 CFTR 104 chaperone proteins 33 charybdotoxin 96, 98 chemokine 79 chlorpropamide 436 CHOP 226, 239–240 CIC-3 140 citrate 63 citric acid cycle 59 CK 19 268 Cl− channels 103 ClC3 103 cleavage of 38 cleave mainly at K⋅R or R⋅R sites 40 CoA pool 60 compound exocytosis 208 conductance 115 confocal microscopy 3 conformation 34 congenital hyperinsulinism of infancy (CHI) 433, 445 connexin proteins 111 connexins 113 connexon 113 consensus model 66 constitutive-like secretion 36 conversion of proinsulin to insulin 34 cooperative kinetics 460 core 41 core–mantle architecture 5 coupling 124 covalent structure of human insulin 32

Subject Index CPE 37 C-peptide 33 C-peptide radioimmunoassay 42 Cre recombinase 247 CREB 230–231, 243, 318, 324 crystal 206 crystallization 204 C-terminal amidation 41 Cx32 118 Cx36 120 Cx36-null mice 124 Cx43 120 CXCR4 275 Cxs 113 cyclin-D 218–231 cyclin-D dependent kinase-4 218 cyclin-dependent kinases 249 cyclins 249 cytokeratin 19 272 cytokine 79–81, 83–84, 222–226, 228–230, 232–234, 238, 240, 243 cytokine NO production 228 cytokines 125, 324 cytoskeletal proteins 119 cytosol 55 cytosolic Ca2+ 123 cytosolic calcium concentration 197 cytotoxic 390

d DAG 162 deﬁnitive endoderm 274 delta 18 delta cells 6 DEND 438, 440–444 dense-core secretory granules 34 depolarization 56 des-31,32 proinsulin 34 des-64,65 proinsulin 34 development 13 developmental delay 438, 441 developmental redirection 285–286, 295, 298, 300–304 diacylglycerol 58 diazoxide 82 dicarboxylate carrier 62 DIDS-sensitive Cl− channels 166 diet induced obesity (DIO) 459

467

differentiation of pancreatic endocrine cells 275 differentiation 13, 267, 322 dihydropyridines DHP 159 directed movements 156 disruption of 40 disulﬁde bonds 32 DNA damage 235, 238 ducts 247 dylipidemia 75, 79–80, 83 dynein 158

e 4E-BP 230–231 E element 20 E-cadherin 124 ECSIT 230, 233 ectopic expression 297, 300 EGF 220 electrical synapses 117 electrical synchronization 121 electrophysiological measurements 148 elements 16 embryoid bodies 273 encoding PC1/3 40 endocrine cell populations 7–8 endocrine glands 118 endocytosis 203 endogenous GK activator(s) 460 endoplasmic stress 390 enteroendocrine 453 enteroendocrine cells 291 Epac 133–138, 141, 143–146 Epac2 135–141, 143–145, 154, 321 epilepsy 438, 441 epsilon cells 4 ER Ca2+ 226 ER stress 42, 75, 81, 324 (ERK1/2) 318 Erk-1/-2 230–232, 242 ER-resident proteins 42 ER-stress 225–226 ES cells 273 exendin-(9–39) 318 exendin-4 276, 318 exocrine 383 exocrine glands 118 exocytosis 57, 61, 196 expression control 460

468

Subject Index

extra-pancreatic cells 303–304 extrapancreatic tissues 285 Extra-Pancreatic Tissues 295

f 5′-ﬂanking 15 factor–basal 16 factor–coactivator 16 factor–factor 16 FADD 230, 233 Fas 75–76, 78–79 fat inﬁltration 383 FFA 58 FFA oxidation 60 ﬁbrils 387 ﬁbroblast growth factor 10 276 ﬁrst phase insulin secretion 327, 384 FLIP 76, 78–79 ﬂuorescence 199, 201 forskolin 133, 138, 141, 142 Foxo1 255 Fred Sanger 31 free fatty acid 80 free fatty acid (FFA) 168 FSH (follicle stimulating hormone) 453 furin 291–292 furin cleavable proinsulin 293 furin-cleavable proinsulin 292 furincleavable 294 fusion pore 166, 203 future perspectives 22

g GABA 104 GABAA [89] and GABAB receptors 163 galanin, somatostatin, and adrenalin 150 gammaaminobutyric acid (GABA) 163 gap junctions 111 gastric-derived G cells 291 gating 116 GEF 231 gene 40, 304 gene therapy 288, 294 genetic disorders 117 genetic engineering 287 genetic pulse-chase 247 ghrelin-producing cells 4 GIP 220, 134, 317

GIRK family 150 GK activator drugs (GKAs) 451 GK regulatory protein (GKRP) 455 GKA effects 458 glibenclamide 436 gliclazide 436 glimepiride 436 glinides 436–437 GLP-1 101, 134, 138, 140, 162, 220, 232, 236, 316, 385 glucagon-like peptide-1 276 glucagon-producing alpha cells 4 gluco-incretins 316 glucokinase 451, 458 glucokinase disease 451 glucolipotoxicity 227, 240–241, 324 glucose competence 327 glucose homeostasis 452 glucose intolerance 325 glucose sensor 451 glucose stimulated insulin secretion (GSIS) 390 glucose 133–134, 138, 142, 145, 320, 451 glucose-regulated promoters 293 glucostat 452 Glut2 promoter 294 glutamate 104 GnRH 454 Golgi apparatus 33 gonadotropes 460 G-protein-coupled receptors 317 Grb2 230–231 growth hormone 219, 229 GSH 64, 67 GSH/GSSG 63 GSK3 230–231 GxTX-1E 101

h H-89 137–138 hematopoietic stem cells 296 hemi-channels 119 heparan sulfate 389 hepatic cell 302 hepatocyte growth factor (HGF) 253, 276 hepatocytes 287, 293, 297–298, 452, 455 “hepato-pancreas” 298 hepatoportal glucose sensor 326

Subject Index HGF 219 high afﬁnity component 154 higher mammalian species 9 highly calcium-sensitive pool (HCSP) 154 HLH 20 home-odomain 18 human 382 human Cx36 gene 126 human embryonic stem (ES) cell 273 human islets 3, 5 hyperglycemia 76, 78–79, 81, 382 hyperinsulinemic hypoglycemia of infancy (PHHI) 456 hyperlipidemic 390 hyperlpasia 219, 221, 236, 243, 385 hypertrophy 221 hypothalamic 453

i IAPP 225–226, 236 IAPP gene 388 iDEND 438, 444 IFNgamma 230, 232–233 IGF-1 219–220, 229–230, 243 IK1 96 IkappaB 233 IkappaBK 233 IL-1 79–80 IL-1ß 225, 228, 230, 232, 234, 241 IL-1beta 75, 78–81 IL-1beta receptor 228, 230, 232, 234, 241 IL-6 225, 228, 230, 232, 234 imaging 196 immature secretory vesicles 33 impaired fasting glucose 384 in vertebrates and are processed similarly 34 incretin 133–134, 138, 142 incretin hormones 290 inﬂammation 79, 225, 228 inﬂammatory stress 225 inner cell mass (ICM) 273 insulin degradation 228 insulin enhancer 15 insulin gene therapy 285–286 insulin granule 134, 138–140, 143–145 insulin mRNA 42 insulin oscillations 126 insulin production 226, 228, 237

469

insulin resistance 215, 221, 224–225, 234, 238–241, 243, 384 insulin secretion 111, 133–135, 137–145, 320 insulin secretory dysfunction 216, 223–224, 228 insulin 13, 31, 43, 147 insulin-like growth factor 1 (IGF1) 276 insulin-positive cells 219 insulin-producing beta cells 4 insulin-treated 384 intercellular communications 111 intercellular junctions 112 intercellular synchronization 124 intermediate cleavage forms 33 intermediate cleavage products 34 interspecies differences 5 intestinal cells 297 intestinal-derived cells 290 intracellular calcium concentrations 9 intra-islet coordination 127 ion channels 91, 149 ionic and metabolic coupling 115 IRAK-1 232 IRS-1 233, 243 IRS-2 227–234, 241–243 Irs2 253, 255 IRS-2 75, 81 islet amyloid 224, 226, 236, 238–240 islet amyloid polypeptide 387 islet amyloid polypeptide (IAPP or amylin) 33 islet architecture 9 islet density 384 islet development and structure 9 islet function 9 islet inﬂammation 79 islet number 386 islet precursors 267 islet size 386 islet transplantation 8, 266 islets 6 islets of Langerhans 452 isoelectric point 34

j JAK2 229–230, 233 Jkn-1/-2 232–233 junctional channels 113

470

Subject Index

k KATP channel 91–92, 134, 139–140, 431–438, 440–441, 443–446 KATP channel, metabolic regulation 435, 437 KATP channel, molecular structure 433 KATP channels 53, 55–56, 61, 66, 75, 82, 149 KCNA5 100 KCNJ11 399, 433, 438, 445 kinesin-1 156 kinesin-3 156 Kir6.1 433 Kir6.2 91, 433–436, 438–445 Kir6.2 (KCNJ11) 92 kiss-and-run exocytosis 166, 207 knock-out mice 117 knockout 325 Kslow 95, 98 KT5720 137 Kv1.5 100 Kv2.1 92, 96, 99, 101, 150 Kv3.2 100 Kv9.3 99 Kv channels 99 Kv-currents 100

l LC-CoA 58, 60–61, 66–67 LDCVs 163 leptin 75, 80, 84 LH (luteinizing hormone) 453 lineage analysis 247 lipid 206 lipid metabolism 58 lipid stress 225 lipolysis 61 lipotoxicity 227, 240–241, 390 liver cells 299 long chain acyl CoA 436 long-chain acyl-CoA 227 low-molecular-weight constituents of the secretory granules 166 loxP 19 L-type Ca2+ channel 91, 94

m macrophage 79, 83–84 macrophages 223, 228 MafA 19 MafB 20 malic enzyme 62–63 malonyl CoA 59–60 map-kinase 231–232, 243 maturing secretory granules 36 maturity onset diabetes of the young (MODY) 433, 456 Mdm2 230–231, 242 mechanism of proteolytic conversion of proinsulin to insulin 39 MEK-1 231 MEKK-1 230, 233 membrane domains 113 membrane fusion 196 membrane potential 148 Menin 255 mesenchymal stem cells 272, 296 metabolic coupling 120 metabolic regulation 53 metabolic stress 225, 228 mice lacking active CPE 41 micro-RNAs 167 microvasculature 196 MIN6 cells 125 mitochondria 59, 75, 81–82 mitochondrial function 392 mitochondrial 54–55 model for Golgi progression 35 MODY 14, 382, 384, 456 monkeys 389 morphometric analysis 7 mouse 6 mouse ES cells 126 mSOS 230–231 mTOR 220, 228, 232, 237, 241–242 multicellular processes 127 multivesicular exocytosis 208 Munc13-1 155 muscle cells 287 muscle weakness 438, 444 muscle-derived cells 291 mutations 43, 117 MyD88 230, 232

Subject Index myosin-5a 158 myosin-6 158

n Na+ channel 91 NADPH 63, 65 NADPH/NADP ratio 67 NADPH/NADP 162 naked plasmid DNA 289, 292 nanometer measurement 200 nateglinide 436 Nav1.3 95 Nav1.7 95 N-CAM 272 necrosis 215, 219, 222, 235, 228, 232, 230, 237, 243 neogenesis 267, 385 neonatal beta-cell growth 224 neonatal diabetes 432–433, 438–443, 446 nestin 272 network of glucose sensors 459 neuroendocrine cells 287, 290 neuroendocrine protein 40 neurosecretory cells 34 NFAT 255 NFkappaB 239 NF-kappaB 75, 81–82 Ngn3+ endocrine progenitors 247 nifedipine 93 NIK 233 nondiabetic obesity 217, 221 nonesteriﬁed fatty acid (NEFAs) 390 non-human primate islets 5 Notch signalling 276 N-terminal prodomains 38 nutritional infertility [13] 455

o O2 consumption 55 ob/ob mice 386 obesity 383 oligomers 390 omega-agatoxin IVA 159 omega-conotoxin GVIA, and SNX482 159 one million islets 384

471

onset 382 optical sections 4 oral agents 384 organoids 126 oscillations 66 oscillatory 56 oxidative stress 225, 227, 235, 239, 241

p 8-pCPT-2′-O-Me-cAMP 136–138, 140, 143 P13K 318 P15ink4b 253 P18ink4c 253 p21 253 p21CIP 230–231 p27 253 P2X prurinergic receptors 119 p300/cAMP 21 p38 230, 232–233, 243 p53 230–231, 242 p57 253 p70S6K 220, 230–232, 242 p90RSK 230–231 pair of basic residues 33 palmitic acid 391 PAM, the peptidyl amidating monooxygenase 41 pancreas 3 pancreatectomy 384 pancreatic beta-cell growth 216, 229, 235–236, 239, 243 pancreatic blood ﬂow 3 pancreatic duct cells 267 pancreatic islets 134–135, 138, 142, 381 pancreatic organogenesis 295 pancreatic progenitor tissue 296 pancreatic sections 7 pancreatic specimens 8 pancreatitis 383 partially 304 pathogenesis of diabetes 223 PAX4 17 Pax4 273 PAX6 17 PC1/3 37, 40 PC2 37, 40 PCD-1 222–223

472

Subject Index

PCs 37–38 PDE 134, 141 PDK-1 18–19, 230–232, 243 Pdx1 273 PDX-1 295–303, 322, 324 PDX-1–VP-16 301–302 peptide-conducting channel 32 PERK 225–226, 239 PERK kinase 225 permanent neonatal diabetes 43 permeability 115 phasing of oscillations 67 phosphatidic acid 58 phosphatidylinositol phosphates 436 phosphofructokinase 57 PI3K 230–232 PI3K pathway 253 piccolo 135, 139, 143, 154 PIP2 436 PIP3 436 pituitary derived cells 289–290 PKA 133–141, 146, 232 PKA inhibitor 134, 137–138, 141, 143 PKB 227–228, 230–232, 241–243 PKB/Akt 227–228, 231–232, 241–243 PKC 225, 227, 230–233, 242 PKCdelta 233 PKCzeta 231 placental lactogen (PL) 219, 221, 253 PNDM 438, 441–442, 444 PNDM (Permanent Neonatal Diabetes Mellitus) 456 polymorphisms 126 porcine 5 posterior foregut 276 post-translational cleavage 32 PP-poor 8 PP-rich 8 prediabetic states 124 Pref-1 229–230 pregnancy 215–218, 221, 224, 237 preproGIP 317 preproglucagon 316 preproinsulin 32 priming/mobilization 155 processing 40, 316 progression 382 prohormone convertases 37 pro-IAPP 40 proinsulin gene 286

proinsulin 32, 34, 40, 294 prolactib 229–230, 233 prolAPP 389 prolatin 219, 221, 229, 230, 233 proliferation 323 properties 34 proprotein 37 protein folding 388 protein kinase C 162 protein synthesis 220, 225, 230–231 protein-thiol reductase 33 Psammomys 76–77 puberty 216 pulsatile insulin secretion 148 pyruvate 62

q 15q14 126 Qa-Qb SNARE SNAP-25 152 Qc-SNARE syntaxin-1a 152

r Raf-1 230–231 Rap1 135, 137, 139, 143, 144 Ras 230–231 Rb 231 readily releasable pool 134, 138, 140 readily releasable pool (RRP) 155 recombinant human insulin from proinsulin 37 recruitment of 123 redox 61, 63 regeneration 326 replication 245 respiration 54, 56–57 retinoic acid 276 retroviral vectors 287 Rim2 135, 139, 143 RIP 230, 233 RIPE3b1 19 rodent 382 rodent islets 3 role in 40 ROS 64–65, 67, 225, 227–228, 241 rough endoplasmic reticulum (RER) 32 R-SNARE VAMP-2 152 ryanodine receptor 94, 139–140

Subject Index s secreted insulin 34 secretory granule proteases 37 secretory granule 41, 94, 147 self-associative properties 34 sequential exocytosis 207 setpoint 452 Shab 98 Shaker 98 Shal 98 Shaw 98 Shc 229–231 signal peptidase 32 signal peptide 32 signal recognition particle (SRP) 32 signal transduction 215, 228–230, 232, 234, 241–242 single beta cells 123 single-chain precursor molecule 31 SK 95 SK channel 96 skeletal muscle 292 SLMVs 163 small-conductance K+ channels 149 SNAP-25 208 SNARE 208 SNARE proteins 152 SOCS 229–230, 233 solubility 34 somatostatin-producing delta cells 4 sonic hedgehog inhibitor 276 spider toxins 101 STAT1 230, 233 STAT3 230, 233 STAT5 229–230, 233 streptozotocin 249 structure 31 structure of human proinsulin 33 succinate 62 sulfonylurea glibenclamide 126 sulfonylureas 75, 82, 385 sulphonylurea receptor (SUR) 433–435, 437, 442–443, 445 sulphonylureas 432 superoxide 64 SUR1 92, 134–135, 139–140, 143, 145, 154, 434–435, 438–440, 442–445 SUR1 (ABCC8) 92 SUR2A 435, 444 SUR2B 435, 444

473

surrogate beta cells 287, 301 surrogate pancreatic beta cells 285 synaptotagmin IX 154 synchronization 112 Synergistic Activation 21 syntaxin 209

t TAB 230, 233 TAK-1 230, 233 TEA 98, 100 tetraethylammonium [TEA] 96 tetrodotoxin (TTX) 95 TGF-beta 253 the Retinoblastoma (Rb) protein 249 the secretory pathway 32 thioredoxin 63–64 tight junction 119 TNDM 438, 441, 444 TNFalpha 222, 225, 228, 230, 232–234 TNGalpha receptor 222, 228, 230, 232–234 TNGalpha 222, 228, 230, 243 tolbutamide 436, 440–441, 444 tomosyn-1 152 TORC-1 230–232 TORC2 230–231 TPPV1 101 TRADD 230, 233 TRAF-2 230, 233 TRAF-6 230, 232–233 trans Golgi network 35 transcription 13 transcription factors 14, 384 transcriptome 119 transdifferentiation 300–303 transgenic 15 transgenic mice 123, 389 transient receptor potential (TRP) cation channel 101 transplantation 5, 266 transport and processing 40 triggering action 148 TRPC 101 TRPC1 103 TRPC4 103 TRPC6 103 TRPM2 103 TRPM4 103

474

Subject Index

TRPM5 103 TRPML 101 TRPV1 103 TRX 64, 67 trypsin 36 T-type Ca2+ channel family (CaV3.1, 3.2, and 3.3) 159 two-photon excitation 199 type 1 382 type 1 diabetes 125 type-1 diabetes 216, 223, 232, 234, 236, 238 type 2 diabetes 126, 316, 381, 433, 436–437, 445–446 type I and type II 37 type-2 diabetes 215–217, 221–229, 232, 234–236, 238–240, 242, 245 tyrosine phosphorylation 229, 231, 243

u UCL 1684 98 UKPDS 436 uncinate process 9 uncouplers 116 uncoupling 123 unfolded protein response or UPR

unfolded protein response 225, 239–240 unitary conductance 120

v variant 117 vectors 288 vesicle 197 vesicle diameter 200 voltage-dependent calcium channels (VDCC) 93, 141 voltage-gated Ca2+ channel: L, R and P/Q 149 voltage-gated K+ channel 98, 150 voltage-gated Na+ channels 94, 149 voltage-gated Na+ currents 95 voltage-sensing K+-channels 65 voltage sensitivity 120

w Wnt3a

42

275

z Z region 17 Zucker diabetic fatty rat 390