TRP Ion Channel Function in Sensory Transduction and Cellular Signaling Cascades (Frontiers in Neuroscience)

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TRP ION CHANNEL FUNCTION in SENSORY TRANSDUCTION and CELLULAR SIGNALING CASCADES Edited by

Wolfgang B. Liedtke Duke University Medical Center Durham, North Carolina

with

Stefan Heller Stanford University School of Medicine Stanford, California

Boca Raton London New York

CRC Press is an imprint of the Taylor & Francis Group, an informa business

FRONTIERS IN NEUROSCIENCE Series Editors Sidney A. Simon, Ph.D. Miguel A.L. Nicolelis, M.D., Ph.D.

Published Titles Apoptosis in Neurobiology Yusuf A. Hannun, M.D., Professor of Biomedical Research and Chairman/Department of Biochemistry and Molecular Biology, Medical University of South Carolina Rose-Mary Boustany, M.D., tenured Associate Professor of Pediatrics and Neurobiology, Duke University Medical Center Methods for Neural Ensemble Recordings Miguel A.L. Nicolelis, M.D., Ph.D., Professor of Neurobiology and Biomedical Engineering, Duke University Medical Center Methods of Behavioral Analysis in Neuroscience Jerry J. Buccafusco, Ph.D., Alzheimer’s Research Center, Professor of Pharmacology and Toxicology, Professor of Psychiatry and Health Behavior, Medical College of Georgia Neural Prostheses for Restoration of Sensory and Motor Function John K. Chapin, Ph.D., Professor of Physiology and Pharmacology, State University of New York Health Science Center Karen A. Moxon, Ph.D., Assistant Professor/School of Biomedical Engineering, Science, and Health Systems, Drexel University Computational Neuroscience: Realistic Modeling for Experimentalists Eric DeSchutter, M.D., Ph.D., Professor/Department of Medicine, University of Antwerp Methods in Pain Research Lawrence Kruger, Ph.D., Professor of Neurobiology (Emeritus), UCLA School of Medicine and Brain Research Institute Motor Neurobiology of the Spinal Cord Timothy C. Cope, Ph.D., Professor of Physiology, Emory University School of Medicine Nicotinic Receptors in the Nervous System Edward D. Levin, Ph.D., Associate Professor/Department of Psychiatry and Pharmacology and Molecular Cancer Biology and Department of Psychiatry and Behavioral Sciences, Duke University School of Medicine Methods in Genomic Neuroscience Helmin R. Chin, Ph.D., Genetics Research Branch, NIMH, NIH Steven O. Moldin, Ph.D, Genetics Research Branch, NIMH, NIH Methods in Chemosensory Research Sidney A. Simon, Ph.D., Professor of Neurobiology, Biomedical Engineering, and Anesthesiology, Duke University Miguel A.L. Nicolelis, M.D., Ph.D., Professor of Neurobiology and Biomedical Engineering, Duke University The Somatosensory System: Deciphering the Brain’s Own Body Image Randall J. Nelson, Ph.D., Professor of Anatomy and Neurobiology, University of Tennessee Health Sciences Center The Superior Colliculus: New Approaches for Studying Sensorimotor Integration William C. Hall, Ph.D., Department of Neuroscience, Duke University Adonis Moschovakis, Ph.D., Institute of Applied and Computational Mathematics, Crete

New Concepts in Cerebral Ischemia Rick C. S. Lin, Ph.D., Professor of Anatomy, University of Mississippi Medical Center DNA Arrays: Technologies and Experimental Strategies Elena Grigorenko, Ph.D., Technology Development Group, Millennium Pharmaceuticals Methods for Alcohol-Related Neuroscience Research Yuan Liu, Ph.D., National Institute of Neurological Disorders and Stroke, National Institutes of Health David M. Lovinger, Ph.D., Laboratory of Integrative Neuroscience, NIAAA In Vivo Optical Imaging of Brain Function Ron Frostig, Ph.D., Associate Professor/Department of Psychobiology, University of California, Irvine Primate Audition: Behavior and Neurobiology Asif A. Ghazanfar, Ph.D., Primate Cognitive Neuroscience Lab, Harvard University Methods in Drug Abuse Research: Cellular and Circuit Level Analyses Dr. Barry D. Waterhouse, Ph.D., MCP-Hahnemann University Functional and Neural Mechanisms of Interval Timing Warren H. Meck, Ph.D., Professor of Psychology, Duke University Biomedical Imaging in Experimental Neuroscience Nick Van Bruggen, Ph.D., Department of Neuroscience Genentech, Inc., South San Francisco Timothy P.L. Roberts, Ph.D., Associate Professor, University of Toronto The Primate Visual System John H. Kaas, Department of Psychology, Vanderbilt University Christine Collins, Department of Psychology, Vanderbilt University Neurosteroid Effects in the Central Nervous System Sheryl S. Smith, Ph.D., Department of Physiology, SUNY Health Science Center Modern Neurosurgery: Clinical Translation of Neuroscience Advances Dennis A. Turner, Department of Surgery, Division of Neurosurgery, Duke University Medical Center Sleep: Circuits and Functions Pierre-Hervé Luoou, Université Claude Bernard Lyon I, Lyon, France Methods in Insect Sensory Neuroscience Thomas A. Christensen, Arizona Research Laboratories, Division of Neurobiology, University of Arizona, Tucson, AZ Motor Cortex in Voluntary Movements Alexa Riehle, INCM-CNRS, Marseille, France Eilon Vaadia, The Hebrew University, Jeruselum, Israel Neural Plasticity in Adult Somatic Sensory-Motor Systems Ford F. Ebner, Vanderbilit University, Nashville, TN Advances in Vagal Afferent Neurobiology Bradley J. Undem, Johns Hopkins Asthma Center, Baltimore, MD Daniel Weinreich, University of Maryland, Baltimore, MD The Dynamic Synapse: Molecular Methods in Ionotropic Receptor Biology Josef T. Kittler, University College, London Stephen J. Moss, University of Pennsylvania Animal Models of Cognitive Impairment Edward D. Levin, Duke University Medical Center, Durham, NC Jerry J. Buccafusco, Medical College of Georgia, Augusta, GA

CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2007 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-10: 0-8493-4048-9 (Hardcover) International Standard Book Number-13: 978-0-8493-4048-2 (Hardcover) This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable eﬀorts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microﬁlming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www. copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-proﬁt organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identiﬁcation and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data TRP ion channel function in sensory transduction and cellular signaling cascades / Wolfgang B. Liedtke, editor. p. ; cm. -- (Frontiers in neuroscience) Includes bibliographical references and index. ISBN-13: 978-0-8493-4048-2 (hardcover : alk. paper) ISBN-10: 0-8493-4048-9 (hardcover : alk. paper) 1. TRP channels. I. Liedtke, Wolfgang B. II. Series: Frontiers in neuroscience (Boca Raton, Fla.) [DNLM: 1. Ion Channels--physiology. 2. Receptors, Sensory--physiology. 3. Signal Transduction--physiology. QU 55.7 T864 2007] QP552.T77T77 2007 572’.3--dc22 Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

2006014151

Series Preface The Frontiers in Neuroscience series presents the insights of experts on emerging experimental techniques and theoretical concepts that are or will be at the vanguard of neuroscience. Books in the series cover topics ranging from methods to investigate apoptosis to modern techniques for neural ensemble recordings in behaving animals. The series also covers new and exciting multidisciplinary areas of brain research, such as computational neuroscience and neuroengineering, and describes breakthroughs in ﬁelds like insect sensory neuroscience, primate audition, and biomedical engineering. The goal is for this series to be the reference that every neuroscientist uses in order to get acquainted with new methodologies in brain research. These books can be given to graduate students and postdoctoral fellows when they are looking for guidance to start a new line of research. Each book is edited by an expert and consists of chapters written by the leaders in a particular ﬁeld. Books are richly illustrated and contain comprehensive bibliographies. Chapters provide substantial background material relevant to the particular subject. Hence, they are not the usual type of method books. They contain detailed ‘‘tricks of the trade’’ and information as to where these methods can be safely applied. In addition, they include information about where to buy equipment and web sites helpful in solving both practical and theoretical problems. Finally, they present detailed discussions of the present knowledge of the ﬁeld and where it should go. We hope that, as the volumes become available, the effort put in by us, the publisher, the book editors, and the individual authors will contribute to the further development of brain research. The extent to which we achieve this goal will be determined by the utility of these books. Sidney A. Simon, Ph.D. Miguel A.L. Nicolelis, M.D., Ph.D. Series Editors

Preface Putting together this volume has been greatly enjoyable! First, our heartfelt gratitude goes to all contributors for making room in their busy professional lives to generate excellent chapters. It takes considerable energy and effort to deliver a complete chapter, which generates no immediate quid pro quo beneﬁt for the chapter author. We are all too aware that this energy and effort could have been invested in the ﬁght-to-survive effort mandatory in 2006, amid narrowing research budgets and increasing workloads with teaching, review commitments, and administrative deadbeat jobs such as animal or internal review board protocols, which stretch our workload toward 14- to 16-hour days. In view of this, again, thank you for contributing highly relevant work. After all, it must be the fascination with TRP channels that drives us—it’s the TRP, stupid! This is highlighted by the number of publications—primary papers and review articles—over time (see Figure P.1), which has currently reached a hyperexponential phase. Where will it go? This fascination has led to a crowding of our ﬁeld, which, in turn, has brought with it some unfortunate consequences. These imply at times undesirable side effects, not unlike those seen in the animal world, when a limited supply is available to a hypermotivated cohort consisting of so many individuals that not all of them are able to satisfy their needs. In a recent example, we learned that an author experienced, after submitting an invited review paper on a TRP-related subject to a scientiﬁc journal (a decent journal, neither a top-tier journal nor an obscure one), a harrowing review process, which consisted of two rounds of ‘‘bean counting’’ with a total of more than 90 points of criticism. Needless to say it was difﬁcult to recognize a signiﬁcant improvement by these modiﬁcations, and the primary submission had received input from several curious and critical readers and had been submitted in technically ﬂawless shape. This leaves room for worry about where our ﬁeld is heading. Is the stress of having to secure an existence as a scientist so overbearing that this leads to a dog-eat-dog attitude? Or is this merely an egregious example of narcissism and allodynic territoriality, which can happen in a rapidly moving ﬁeld, and will remain, hopefully, an exception? This volume has been edited carefully, yet one major editorial principle has been to let authors have their say without the constraints imposed by detail-obsessed reviewers. Because the ﬁeld of TRP channels is new and rapidly moving, this also means that some “hot topic” areas have been covered by more than one author, and the viewpoints of these authors have not been edited for mutual alignment. Time will show where we will be taken. This volume does not claim to represent an all-comprehensive view of the ﬁeld of TRP channel biology (for this, the interested reader is referred to numerous excellent review papers referenced in the individual chapters; the latest wave of them is in an extra section of Annual Review of Physiology, vol. 68, pp. 619–736,

600 number of papers

At least 4k papers since 1969! 400 original papers reviews

200

0 1980

1985

1990

1995

2000

2005

year of publication

FIGURE P.1 Number of TRP papers. (Courtesy of Bernd Nilius, Leuven, Belgium.)

2006). This book, however, does shed light on selected topics of outstanding interest in the TRP arena, and the spotlight is cast by individuals who did earn their mettle. A book on a hyper-rapidly moving topic such as TRP channels does raise the question of how such a topic can be dealt with in the publishing world. Web-based follow-up editions represent one hypothetical tool in response to this challenge, which we deem an attractive possibility. Wolfgang B. Liedtke and Stefan Heller

The Editors Wolfgang B. Liedtke, MD, PhD, is an assistant professor at the Duke University Center for Translational Neuroscience, a joint venture between the Department of Neurobiology and the Department of Medicine/Division of Neurology at Duke. He is a board-certiﬁed neurologist who provides part-time clinical service in an outpatient pain clinic at Duke University Medical Center. Most of his effort is dedicated toward his laboratory, which focuses on molecular mechanisms of neurosensory transduction, in particular by TRP ion channels. Dr. Liedtke is well accomplished in this ﬁeld, and he was awarded a Klingenstein Fellowship in Neuroscience in 2004. He went through residency training in his native Germany in neurology and psychiatry and followed up with a neuropathology fellowship at Albert Einstein College of Medicine in New York City, where he held a Feodor Lynen Fellowship of the Alexander von Humboldt Foundation, Bonn, Germany (his mentor was Dr. Cedric S. Raine). Next, remaining in New York City, he learned molecular biology “from scratch” at The Rockefeller University (with Dr. Jeffrey M. Friedman). At Rockefeller University, he met Dr. Stefan Heller, which led to a highly successful collaboration. In 2004, Dr. Liedtke assumed a tenure-track faculty position at Duke University in Durham, North Carolina. Stefan Heller, PhD, recently joined the faculty of Stanford University School of Medicine as an associate professor of otolaryngology—head and neck surgery and an associate professor of molecular and cellular physiology. His initial faculty appointment was at Harvard Medical School, where he was assistant professor for otolaryngology in 2000. In 2005, he was appointed as the James Wiggins associate professor of otolaryngology with courtesy appointments in the Program in Neuroscience and the Harvard–MIT Division of Health Sciences Technology. Dr. Heller’s group focuses on elucidating the molecular principles of mechanoreception by TRP ion channels. This work is a continuation of his collaboration with Dr. Wolfgang B. Liedtke, which started in New York City in the ﬁnal years of a postdoctoral fellowship with Dr. A. James Hudspeth. In addition, Dr. Heller has distinguished himself as one of the preeminent authorities on stem cells in the inner ear. His laboratory is ﬁnding solutions for cell replacement in the damaged cochlea using stem cells and progenitor cells. Dr. Heller’s honors include receiving the McKnight Neuroscience of Brain Disorders Award and awards from the Deafness Research Foundation, the National Organization for Hearing Research, the March of Dimes Birth Defects Foundation, the American Neurotology Society, and the Albert and Ellen Grass Foundation.

Contributors Joel Abramowitz National Institute of Environmental Health Sciences Research Triangle Park, North Carolina Silvia Amadesi Departments of Surgery and Physiology University of California San Francisco, California Yaniré N. Andrade Grup de Canalopaties Universitat Pompeu Fabra Barcelona, Spain Maite Arniges Grup de Canalopaties Universitat Pompeu Fabra Barcelona, Spain Maureen M. Barr School of Pharmacy University of Wisconsin at Madison Madison, Wisconsin Lutz Birnbaumer National Institute of Environmental Health Sciences Research Triangle Park, North Carolina Peter M. Blumberg Laboratory of Cellular Carcinogenesis and Tumor Promotion National Cancer Institute Bethesda, Maryland

Sebastian Brauchi Universidad Austral de Chile Valdivia, Chile and Centro de Estudios Cientiﬁcos Valdivia, Chile Joseph E. Brayden Department of Pharmacology University of Vermont College of Medicine Burlington, Vermont Nigel W. Bunnett Departments of Surgery and Physiology University of California San Francisco, California David M. Cohen Division of Nephrology and Hypertension Oregon Health & Science University and the Portland Veterans Affairs Medical Center Portland, Oregon Kai Cui Institute of Neuroscience Shanghai Institutes for Biological Sciences Chinese Academy of Sciences Shanghai, China Patrick Delmas Laboratoire de Neurophysiologie Cellulaire, CNRS Marseille, France

Anne Duggan Departments of Anesthesiology, Physiology and Neurology Northwestern University Institute for Neuroscience Chicago, Illinois Catherine Dulac Department of Molecular and Cellular Biology Howard Hughes Medical Institute Harvard University Cambridge, Massachusetts Scott Earley Department of Pharmacology University of Vermont College of Medicine Burlington, Vermont Petra Eder Institute of Pharmaceutical Sciences, Pharmacology and Toxicology Karl Franzens University of Graz Graz, Austria Jacqueline Fernandes Grup de Canalopaties Universitat Pompeu Fabra Barcelona, Spain Antonio Ferrer-Montiel Instituto de Biología Molecular y Celular Universidad Miguel Hernández Alicante, Spain Jaime García-Añoveros Departments of Anesthesiology, Physiology and Neurology Northwestern University Institute for Neuroscience Chicago, Illinois

Rachelle Gaudet Department of Molecular and Cellular Biology Harvard University Cambridge, Massachusetts Aurélie Giamarchi Laboratoire de Neurophysiologie Cellulaire, CNRS Marseille, France Andrew Grant Departments of Surgery and Physiology University of California San Francisco, California Klaus Groschner Institute of Pharmaceutical Sciences, Pharmacology and Toxicology Karl Franzens University of Graz Graz, Austria Marilia Z. P. Guimaraes Departamento de Farmacologia Basica e Clinica Universidade Federal do Rio de Janeiro Rio de Janeiro, Brazil Christian Harteneck Institut für Pharmakologie Charité Campus Benjamin Franklin Berlin, Germany Stefan Heller Departments of Otolaryngology—HNS and Molecular & Cellular Physiology Stanford University School of Medicine Stanford, California

Joachim Hoyer Department of Internal MedicineNephrology Philipps University Marburg, Germany

Emily R. Liman Department of Biological Sciences/Neurobiology University of Southern California Los Angeles, California

Yuji Imaizumi Department of Molecular and Cellular Pharmacology Nagoya City University Nagoya, Japan

Ivan M. Lorenzo Grup de Canalopaties Universitat Pompeu Fabra Barcelona, Spain

Sven-Eric Jordt Department of Pharmacology Yale University School of Medicine New Haven, Connecticut Changsoo Kim School of Biological Sciences Chonnam National University Gwangju-Si, Korea

David D. McKemy Department of Biological Sciences Neurobiology Section and School of Dentistry University of Southern California Los Angeles, California Katsuhiko Muraki Laboratory of Cellular Pharmacology Aichi Gakuin University Nagoya, Japan

Ralf Köhler Department of Internal MedicineNephrology Philipps University Marburg, Germany

Masahiro Nagasawa Institute for Molecular & Cellular Regulation Gunma University Maebashi, Japan

Itaru Kojima Institute for Molecular & Cellular Regulation Gunma University Maebashi, Japan

Roger G. O’Neil Department of Integrative Biology and Pharmacology The University of Texas Health Science Center Houston, Texas

Ramon Latorre Centro de Estudios Cientiﬁcos Valdivia, Chile Wolfgang B. Liedtke Center for Translational Neuroscience Duke University Medical Center Durham, North Carolina

Gerardo Orta Centro de Estudios Cientiﬁcos Valdivia, Chile Rosa Planells-Cases Centro de Investigación Príncipe Felipe Valencia, Spain

Tim D. Plant Institute for Pharmacology and Toxicology Philipps University Marburg, Germany James W. Putney, Jr. National Institute of Environmental Health Sciences Research Triangle Park, North Carolina Stacey Reading Department of Pharmacology University of Vermont College of Medicine Burlington, Vermont Christoph Romanin Institute of Pharmaceutical Sciences, Pharmacology and Toxicology Karl Franzens University of Graz Graz, Austria Tamara Rosenbaum Departamento de Bioﬁsica Instituto de Fisiología Celular Universidad Nacional Autónoma de México México City, México Rainer Schindl Institute of Pharmaceutical Sciences, Pharmacology and Toxicology Karl Franzens University of Graz Graz, Austria

Sidney A. Simon Department of Neurobiology Duke University Durham, North Carolina Rainer Strotmann Institut für Biochemie Abteilung Molekulare Biochemie Universität Leipzig Leipzig, Germany Arpad Szallasi Department of Pathology Monmouth Medical Center Long Branch, New Jersey and Department of Pathology Drexel University College of Medicine Philadelphia, Pennsylvania Ji Ying Sze Department of Anatomy and Neurobiology College of Medicine University of California Irvine, California Makoto Tominaga Section of Cell Signaling Okazaki Institute for Integrative Bioscience National Institutes of Natural Sciences Okazaki, Japan

Günter Schultz Institut für Pharmakologie Charité Campus Benjamin Franklin Berlin, Germany

W. Daniel Tracey, Jr. Departments of Anesthesiology, Cell Biology and Neurobiology Duke University Medical Center Durham, North Carolina

Munekazu Shigekawa Department of Human Life Sciences Senri Kinran University Osaka, Japan

Miguel A. Valverde Grup de Canalopaties Universitat Pompeu Fabra Barcelona, Spain

Guillermo Vargas Centro de Estudios Cientiﬁcos Valdivia, Chile

Eda Yildirim National Institute of Environmental Health Sciences Research Triangle Park, North Carolina

X. Z. Shawn Xu Life Sciences Institute Department of Molecular and Integrative Physiology University of Michigan Ann Arbor, Michigan

Xiao-bing Yuan Institute of Neuroscience Shanghai Institutes for Biological Sciences Chinese Academy of Sciences Shanghai, China

Abstract TRP ion channels were ﬁrst described in Drosophila melanogaster in 1989 and in mammals several years later. In 1997, TRPV1, a member of the TRP channel superfamily (now with more than 60 members in vertebrates and invertebrates but not in bacteria and plants), was described to respond to the pungent ingredients of hot pepper, then named capsaicin receptor. Ever since we have witnessed an explosion of activity in this ﬁeld of scientiﬁc inquiry for obvious reasons. TRP ion channels are critical elements in signal transduction of cellular signaling cascades and of neurosensory processes, which are involved in all ﬁve senses. This book, TRP Ion Channel Function in Sensory Transduction and Cellular Signaling Cascades presents 31 chapters written by researchers who have made these key discoveries, such as Dr. Lutz Birnbaumer who discovered mammalian TRP channels, and who continues to conduct TRP ion channel research at the cutting edge of this hyperdynamic area. Because of the burgeoning nature of the ﬁeld, this book does not represent an all-comprehensive view on TRP channel biology. However, it does shed light on selected topics of outstanding interest in the TRP arena, such as signal transduction in axonal pathﬁnding, and vascular, renal, auditory, and nociceptive functioning, to name a few, and the spotlight is cast by an international cast of outstanding chapter authors.

Contents Chapter 1

The TRPC Family of Ion Channels: Relation to the TRP Superfamily and Role in Receptor- and Store-Operated Calcium Entry.............................................................1

Joel Abramowitz, Eda Yildirim, and Lutz Birnbaumer Chapter 2

Multiple Mechanisms of TRPC Activation .......................................31

James W. Putney, Jr. Chapter 3

TRPC2 and the Molecular Biology of Pheromone Detection in Mammals ............................................................................................45

Emily R. Liman and Catherine Dulac Chapter 4

TRP Channels and Axon Pathﬁnding................................................55

Kai Cui and Xiao-bing Yuan Chapter 5

TRPV1 Receptors and Signal Transduction......................................69

Tamara Rosenbaum and Sidney A. Simon Chapter 6

Complex Regulation of TRPV1 by Vanilloids ..................................85

Arpad Szallasi and Peter M. Blumberg Chapter 7

TRPV2: A Calcium-Permeable Cation Channel Regulated by Insulin-Like Growth Factors.......................105

Itaru Kojima and Masahiro Nagasawa Chapter 8

Molecular Mechanisms of TRPV4 Gating......................................113

Stefan Heller and Roger G. O’Neil Chapter 9

TRPV4: A Multifunctional Nonselective Cation Channel with Complex Regulation..................................................125

Tim D. Plant and Rainer Strotmann

Chapter 10 TRPV4 and TRPM3 as Volume-Regulated Cation Channels .........141 Christian Harteneck and Günter Schultz Chapter 11 TRPA1: A Sensory Channel of Many Talents.................................151 Marilia Z. P. Guimaraes and Sven-Eric Jordt Chapter 12 TRPA1 in Auditory and Nociceptive Organs ..................................163 Jaime García-Añoveros and Anne Duggan Chapter 13 TRPM8: The Cold and Menthol Receptor ......................................177 David D. McKemy Chapter 14 Activation Mechanisms and Functional Roles of TRPP2 Cation Channels..............................................................189 Aurélie Giamarchi and Patrick Delmas Chapter 15 The Ca2+-Activated TRP Channels: TRPM4 and TRPM5..............203 Emily R. Liman Chapter 16 Genetics Can Be Painless: Molecular Genetic Analysis of Nociception in Drosophila...........................................213 W. Daniel Tracey, Jr. Chapter 17 TRPV Family Ion Channels and Other Molecular Components Required for Hearing and Proprioception in Drosophila ...................................................................................227 Changsoo Kim Chapter 18 The TRPV Channel in C. elegans Serotonergic Neurons...............243 Ji Ying Sze Chapter 19 TRP Channel Functioning in Mating and Fertilization ..................257 X. Z. Shawn Xu and Maureen M. Barr Chapter 20 The Role of TRP Channels in Thermosensation.............................271 Makoto Tominaga

Chapter 21 Voltage and Temperature Gating of ThermoTRP Channels ...........287 Ramon Latorre, Guillermo Vargas, Gerardo Orta, and Sebastian Brauchi Chapter 22 TRPV Channels’ Function in Osmo- and Mechanotransduction .......................................................................303 Wolfgang B. Liedtke Chapter 23 TRP Channel Trafﬁcking .................................................................319 Rosa Planells-Cases and Antonio Ferrer-Montiel Chapter 24 Protein–Protein Interactions in TRPC Channel Complexes ...........331 Petra Eder, Rainer Schindl, Christoph Romanin, and Klaus Groschner Chapter 25 Structural Insights into the Function of TRP Channels ..................349 Rachelle Gaudet Chapter 26 Functional Signiﬁcance of Transient Receptor Potential Channels in Vascular Function.........................................361 Scott Earley, Stacey Reading, and Joseph E. Brayden Chapter 27 Role of TRPV4 in the Mechanotransduction of Shear Stress in Endothelial Cells................................................377 Ralf Köhler and Joachim Hoyer Chapter 28 A New Insight into the Function of TRPV2 in Circulatory Organs.......................................................................389 Katsuhiko Muraki, Munekazu Shigekawa, and Yuji Imaizumi Chapter 29 The Role of TRPV4 in the Kidney .................................................397 David M. Cohen Chapter 30 The TRPV4 Channel in Ciliated Epithelia......................................413 Yaniré N. Andrade, Jacqueline Fernandes, Ivan M. Lorenzo, Maite Arniges, and Miguel A. Valverde

Chapter 31 Protease-Activated Receptors: Mechanisms by Which Proteases Sensitize TRPV Channels to Induce Neurogenic Inﬂammation and Pain ...............................421 Andrew Grant, Silvia Amadesi, and Nigel W. Bunnett Index......................................................................................................................441

1

The TRPC Family of Ion Channels: Relation to the TRP Superfamily and Role in Receptor- and StoreOperated Calcium Entry Joel Abramowitz Eda Yildirim Lutz Birnbaumer National Institute of Environmental Health Sciences

CONTENTS TRP Channels: Diversity of Form and Function ......................................................2 TRP Superfamily Genes and Their Gene Products ..................................................4 TRP Subfamilies............................................................................................4 Genomic Structures .....................................................................................11 Direct Chromosomal Repeats......................................................................12 Mechanism(s) of TRPC Activation .........................................................................14 Receptor-Mediated Activation of TRPCs Requires Phospholipase C Activation: A Role for PDZ Scaffolds? ..........................14 Activation by Conformational Coupling: The Role for IP3 Receptor as an Activator of TRPC through Protein-Protein Interaction ..........................................................................15 Activation of TRPCs by Diacylglycerol (DAG) and Inhibition of TRPCs by Protein Kinase C (PKC) .......................................................17 Activation of Ca2+ Entry by Channel Translocation from Endomembranes to the Plasma Membrane........................................17 Physiological Role for TRPC Channels as Electrogenic Devices That Couple GPCR-Gq Activation to Voltage-Gated Ca2+ Channel Activation ..............................................................................18 A Role for Tyrosine Kinases in Voltage-Gated Ca2+ Channel-Independent Ca2+ Inﬂux ................................................................19

1

2

TRP Ion Channel Function in Sensory Transduction

Studies on the Role(s) of Tyrosine Phosphorylation in TRPC Function ...............19 TRPCs and SOCE .......................................................................................23 Acknowledgments....................................................................................................24 References................................................................................................................24

TRP CHANNELS: DIVERSITY OF FORM AND FUNCTION The Drosophila trp mutation is responsible for the phenotype called transient receptor potential, an alteration of the ﬂy’s electrorentinogram in which its sustained phase is missing.1,2 The responsible gene was cloned in 1989.3 Its amino acid sequence predicted a protein with eight hydrophobic segments that could potentially form transmembrane segments. Puriﬁcation and cloning of a calmodulin-binding protein from Drosophila heads showed it to be a homologue of trp. It received the name trp-like or trpl.4 Its discoverers highlighted the existence of limited sequence similarities between trp/trpl and voltage-sensitive Na+ and Ca2+ channels. Expression of trpl in silkworm cells of Spodoptera frugiperda (Sf9 cells) did indeed lead to appearance of cation channels.5 In keeping with both a role for trp and trpl in insect phototransduction, and the fact that insect phototransduction is biochemically akin to mammalian signal transduction based on the Gq-PLC pathway instead of a transducin-phosphodiesterase (Gt-PDE) pathway,6,7 the trpl channels expressed in Sf9 cells could be activated by a Gq-coupled GPCR.8 Activation of the Gq-PLC signaling system results in hydrolysis of PIP2 with formation of the second messengers diacylglycerol (DAG) and inositol trisphosphate (IP3), followed by IP3-induced release of Ca2+ from intracellular stores and the activation of type-C protein kinases (PKCs) by the combined action of DAG and the released Ca2+. Further, the depletion of intracellular Ca2+ stores then activates Ca2+-permeable cation channels in the plasma membrane. The molecular basis by which store depletion activates plasma membrane Ca2+ entry channels is as yet incompletely deﬁned. Two questions need to be answered: (1) which molecules make up the channels that mediate the store depletion–activated Ca2+ entry?; and (2) by what mechanism do the stores ‘‘inform’’ the plasma membrane channels of their state of replenishment? TRP channels have been postulated as the pore-forming molecules through which store depletion–activated Ca2+ entry takes place.9 While there are data in support of this hypothesis, the ﬁnal word is not in, and much is still to be elucidated (vide infra The ROCE SOCE Conundrum). On the other hand, recent studies that screened for a Ca2+ sensor have successfully identiﬁed a single pass membrane protein termed STIM1 as the store’s Ca2+ sensor.10,11 Interestingly, store depletion promotes a translocation of STIM1 from endomembranes to the plasma membrane,12 where it presumably ‘‘talks’’ to the Ca2+ entry channels. Ca2+ entering after activation of the PLC-IP3R store depletion pathway serves both as a substrate for the sarcoplasmic-endoplasmic reticulum Ca2+ pumps (SERCAs), which replenish the depleted stores, and as a signaling molecule. We will refer to Ca2+ entry that follows activation of PLC by receptors as receptor-operated Ca2+ entry or ROCE. Store depletion can also be caused by inactivating SERCA pumps with an inhibitor such as thapsigargin13,14 or by loading cells with Ca2+

The TRPC Family of Ion Channels

3

chelator that acts as a sink for Ca2+ that leaks passively from the stores.15 Both maneuvers activate Ca2+ entry that can be assessed either with ﬂuorescent indicator dyes, such as fura2, in which case it is referred to as capacitative Ca2+ entry (CCE16,17), or as store-operated Ca2+ entry (SOCE) or by electrophysiologic means, where Ca2+ entry presents itself as an inward Ca2+ current, termed Ca2+ releaseactivated Ca2+ current (Icrac).15,18 Icrac and CCE are commonly accepted as being the measures of the same phenomenon. In 1992, Hardie and Minke19 showed that the missing sustained phase of the electroretinograms of trp mutant Drosophila eyes has as its underlying basis the absence of a Ca2+ conductance. In 1993, they formally raised the question whether the trp and trpl proteins might be functional homologues of capacitative Ca2+ entry channels, and by extension the pore-forming molecules of Icrac channels20 in mammalian cells. The ﬁnding that a trpl/trp chimera could be activated by store depletion in Sf9 cells21 lent strong support to Hardie and Minke’s hypothesis. The mammalian homologues of Drosophila trp genes (TRPs) were cloned to test Hardie and Minke’s hypothesis. Six such homologues were identiﬁed in our initial 1995–96 screen,22–24 and a seventh was discovered three years later.25 Initially called TRPs, they are now referred to as TRPCs.26 Expression and assembly studies have shown that TRPCs can selectively form heteromeric complexes, such as 1:2, 1:3, 1:5, 4:5, 3:6:7 and 1:4:5 that co-immunoprecipitate.27–29 Presumably in all cases active channels are tetrameric. Unexpectedly, independent research from several laboratories uncovered the existence of TRP-related cation channels that together constitute a ‘‘superfamily’’ whose members play differing and sometimes still unknown roles in cellular physiology. One set of TRP-related channels, with a role in pain and thermosensing, was uncovered in David Julius’s laboratory. In 1997, Julius isolated the cDNA that encodes the capsaicin (also vanilloid) receptor (VR1) through expression cloning.30 This was followed, shortly afterward, by the cloning of its close relative VRL1.31 Both turned out to be heat sensors and structural homologues of the ﬂy trp channels. Independently, VRL1 was also cloned in 1999 as a growth factor–activated channel, which translocated from endomembranes to the plasma membrane.32 Between 1999 and 2000, two groups, one in the Netherlands and one in Boston, identiﬁed two epithelial calcium transporters: renal ECaC and intestinal CaT. Amino acid sequence analysis revealed ECaC (also CaT2) and CaT (also ECaC2) to be related to VR1 and VRL1, and hence to TRPs. In 2001, the original mammalian TRPs were renamed to TRPCs (for classic or canonical) and VRs and ECaC/CaT were renamed to TRPVs, of which there are six. Mlsn1 (melastatin 1), identiﬁed in 1998 as a gene product down-regulated in melanoma cells,33 is the founding member of the TRPMs, of which there are eight. As these families were being identiﬁed, it became evident that there existed other, more distant relatives that include the polycystins34 and their relatives, TRPPs 1–4 (reviewed in reference 35). Genes responsible for mucolipidosis also encode TRPrelated proteins.36 At present we recognize three TRPMLs. A mechanosensory transduction channel, TRPA1, is the latest addition to mammalian TRP genes.37,38 TRPA1’s most outstanding structural characteristic is an unusually large number of ankyrin motifs in its N-terminus.

4

TRP Ion Channel Function in Sensory Transduction

Most TRPs are calcium-permeable nonselective cation channels, with notable exceptions: ECaC/CaT channels are highly selective for Ca2+,39–41 and TRPM4/5 are nonselective monovalent cation channels activated by Ca2+ without being permeant to Ca2+ (reviewed in reference 42). TRPM2 and TRPM6/7 incorporate enzymatic functions into their C-termini. TRPM2 has a NUDIX domain able to bind ADP-ribose and to sense H2O2; TRPM6/7 carry an atypical (alpha) protein kinase domain; and while TRPV1, 2, and 3 sense and are activated by distinct temperatures, TRPM8 (also CMR, for cold and menthol receptor) is activated upon cooling.43 TRPV4 and TRPM3 are osmo-sensitive. Physiologically, TRPV4 participates in mediating pain sensations, and TRPM5 is a taste transduction channel expressed in sensory neurons mediating bitter, sweet, and amino acid (unami) tastes. TRPC3 has been proposed to be the melanopsin-activated transduction channel of the intrinsically photosensitive retinal ganglion cells (ipRGCs) responsible for entrainment of the circadian clock of the suprachiasmatic nucleus.44,45 TRPC3 and TRPC6 were recently implicated as essential components of the machinery guiding Ca2+-dependent growth cone turning of pontine neuron axon extensions,45 and Xenopus TRPC1 was identiﬁed by Wang and Poo46 in a similar phenomenon whereby netrin guides axonal growth of Xenopus spinal neurons. In a parallel study, TRPC3 and TRPC6 were implicated in BDNF-directed axonal outgrowth from cerebellar granule cells.47 In rodents, TRPC2 has been shown to be the transduction channel activated in vomeronasal sensory neurons in response to activation of vomeronasal pheromoneresponsive GPCRs48,49 (reviewed in references 50 and 51). Recently, TRPA1, also ANKTM1, a channel with fourteen N-terminal ankyrin repeats, was characterized as a channel that transduces noxious cold sensation as well as the mechanical bending of stereocilia of inner ear hair cells.37,38

TRP SUPERFAMILY GENES AND THEIR GENE PRODUCTS TRP SUBFAMILIES As is evident from the above discussion, a variety of different approaches involving many laboratories—some working in parallel—were involved in identifying the various TRP channels and hence naming the cloned genes in a somewhat unruly manner. The nomenclature of the C, M, and V channels has been agreed upon.26 The use of TRPP for the PKD family of channels is still tentative. Table 1.1 tabulates the TRP genes we analyzed and provides their mRNA accession numbers; the lengths, in amino acids, predicted by their coding sequences (CDS), restricted for the most part to the longest if splice variants exist; location of the TRP channel domain (ion channel domain) within the cDNAs; and the genomic accession numbers, chromosomal loci, and number of exons, as predicted by GenBank’s annotated genomic contigs. Using the ion channel domains identiﬁed in Table 1.1 as a basis, we constructed a phylogenetic tree that in turn organized members of the C-, V-, M-, P-, ML-, and A-type TRP channels into subgroups (Figure 1.1). The years in which their founding members were identiﬁed are highlighted.

TRPM1 TRPM2 TRPM3 TRPM4 TRPM5 TRPM6

TRPC7

TRPC6

TRPC5

TRPC4

TRPC3

TRPC2

TRPC1

TRP Channel

Human Human Human Human Human Human

Murine Human Murine Human Murine Human Murine Human Murine Human Murine

Human Murine Human

Species

AF071787 AB001535 AJ505025 AF497623 AF177473 AF448232

U31110 U73625 Pseudo X89067 AF11108 U47050 AF190645 AF175406 AF011543 AF054568 AF029983 AF080394 U49069 AJ272034 AF139923

mRNA Accession Number

1533 1503 1707 1214 1165 2022

1172 848 836 977 974 973 975 931 930 862 862

793 809

# of AA

720-1120 701-1100 769-1174 430-920 581-1020 669-1133

520-970 310-690 310-690 280-640 280-640 280-650 280-650 310-750 310-750 310-690 310-690

300-714 300-714

Ion Channel Domain

NT_010363 NT_011515 NT_008580 NT_011109 NT_033238 NT_008580

NT_005832 NW_000355 NT_035090 NT_033927 NW_000328 NT_016354 NT_035972 NT_033922 NW_000186 NT_025319 NW_042636 NT_009151 NW_000350 NT_037664 NW_000081

Genome Accession Number

C 2085178-2185564 1085877-1175698 C 2314499-2399547 11792968-11847007 C 573099-591648 C 6501941-6667540

3300676-3384487 C 6496700-6541991 63880-74537 2272990-2336104 C 19937673-19937815 C 11488244-11542672 C 136113-205720 C 6785134-7018278 C 3416035-3579378 C 2989179-3297641 C 3233132-3540959 C 4866094-4998380 C 5510957-5648851 C 424567-568641 C 2431698-2555118

Gene Locus in Contig/Gene

15q13-q14 21q22.3 9q21.11 19q13.32 11p15.5 9q21.13

3q22-q24 9 51.0 cM 11p15.3-p15.4 11 q13.2 7 50.0 cM 4q27 3 18.2 cM 13q13.1-q13.2 3 28.6 cM Xq23 X 63.0 cM 11q21-q22 9 1.0 cM 5q31.1 13 B2

Locus

(continued)

27 32 11 29 24 39

21 11 11 13 13 10 10 13 13 11 11

13 13

# of Exons

TABLE 1.1 TRP Channel Superfamily GenBank Accession Numbers and Numerical Parameters of Their mRNAs and Genes

The TRPC Family of Ion Channels 5

Human Human

Human Human Human Human Human Human

Human

Human Human Human Human

Human Human Human

Human

TRPM7 TRPM8

TRPV1 TRPV2 TRPV3 TRPV4 TRPV5 TRPV6

TRPP1

TRPP2 TRPP3 TRPP4 TRPP5

TRPML1 TRPML2 TRPML3

TRPA1

Y10601

AF287269 AY083533 AF475085

L39891 U24497* U50928 AF073481 AF116458 AF118125

AY131289 AJ487963 AY118268 AJ296305 AJ271207 AF365927

AF346629 AY090109

mRNA Accession Number

1119

580 540 553

3638 4303 968 805 2253 609

839 764 791 871 729 725

1864 1104

# of AA

796-959

354-490 342-506 376-501

2301-3400 198-705 78-585 1707-2171 33-489

401-743 365-704 414-710 444-750 301-600 301-600

683-1158 626-1043

Ion Channel Domain

NT_008183.18

NT_077812.2 NT_032977.7 NC_000001

L39891 NT_010552 NT_006204 NT_030059 NT_011523 NT_016714

NT_010692 NT_010718 NT_010692 NT_009770 NT_007914 NT_007914

NT_010194 NT_005120

Genome Accession Number

C24841572-24786439

191108-203260 39282589-39210261 C85226190-85195787

3648-51866 C 663007-694871 7643751-7713856 C 7312540-7354872 C 163711-171370 365617-420393

C 2321912-2365875 9021374-9042801 C 2269657-2314460 C 790403-840721 C 3218677-3244315 C 3182370-3196917

C 5824294-5950796 756514-858635

Gene Locus in Contig/Gene

*TRPP1 accession number used for multiple sequence alignment analysis. Adapted from Birnbaumer et al.35

Species

TRP Channel

8q13

19p13.3-p13.2 1p22 1p22.3

4q21-23 10q24 22q13.31 5q31

16p13.3

17p13.3 17p11.2 17p13.3 12q23-q24.1 7q35 7q33-q34

15q21 2q37.1

Locus

27

13 13 13

50 23 15 16 1 14

17 15 18 15 15 15

39 25

# of Exons

TABLE 1.1 (Continued) TRP Channel Superfamily GenBank Accession Numbers and Numerical Parameters of Their mRNAs and Genes

6 TRP Ion Channel Function in Sensory Transduction

The TRPC Family of Ion Channels

7

FIGURE 1.1 (Left) Phylogenetic tree illustrating the successive discovery of TRP and TRPrelated channel. Years denote when the founding member of each subfamily was identiﬁed. (Right) Functional diversity among the major subfamilies of TRP channels. The ﬁgure highlights not only the diversity of channels but also the diversity of the regulatory signals impinging on their function. Note in the left panel that several TRP-related channels were independently discovered by more than one group, leading to multiple GenBank accession numbers and to confusing names. This was resolved in 2002, as shown with the C, V, and M nomenclature in the right panel.26

Figure 1.2 presents an amino acid alignment in which we compare the ion channel domains of a representative member of each of the six families (C, V, M, P, ML, A), highlighting in white on black background positions at which three or more amino acid identities occur among the six sequences. The result documents the “relatedness” of the six subfamilies. The segregation of TRP channels into the C, M, V, P, ML, and A subfamilies emerges very clearly from these alignments. Kyte and Doolittle analysis and subsequent glycosylation scanning mutagenesis and limited epitope mapping led to the deﬁnition of six transmembrane domains in TRPC3 preceded by a cytosolic hydrophobic domain (h or h1).52 Based on both the individual hydrophobicity plots and TRPC3 as a template, we deﬁned the putative transmembrane domains of all the TRPC channels listed in Table 1.1. We used the annotations that accompany the genomic sequence ﬁles to reconstruct the open reading frames (ORFs) of the C-, V-, M-, P-, ML-, and A-type TRP mRNAs

8

TRP Ion Channel Function in Sensory Transduction TRPP2

GL WGT R L ME E S S T N R E K Y L K S V L R E L V T Y L L F L I V L C I L T Y GMMS S N V Y Y Y T R MMS QL F L

TRPP2

D T P V S K T E K T N F K T L S S ME D F WK F T E GS L L D GL Y WK M QP S N QT E A D N R S F I F Y E N L L L G V

TRPC3 TRPM2 TRPP2

- - - - - - - - - - - - - - - - - - - - - - L A I K Y E V K - - K F V A H P N CQQQL L - - - A QK L L T R V S E A WG K T T C L Q L A L - - E A K D MK F V S H GG I - QA F L P R I RQL RV RN GS CS I P QDL RDE I K E CY D V Y S V S S E D R A P F GP RNG

- I WY E N L S - - K V WWGQL S V D N A WI Y T S E K D L N

TRPC3 TRPM2 TRPP2 TRPV1

RE Q R- H WG - - -

T I -

S A A -

TRPC3 TRPM2 TRPP2 TRPV1

I V L PN

C R

- L G HL N I VVR HDM

TRPC3 TRPM2 TRPP2 TRPA1 TRPML1 TRPV1

E E -

K R -

EL QL - I - - - A

A L D - -

T S -

DR - TG NR

F F G L

E V L

GI - L - - -

R - - D CGE I L E - - - - - - - - DGL P P

I F

- - - L MK RI H - - - - - K ME

- AA L R- TG

- I L QL L SDF H RSF N - - - - - - - - - - - D RVT GE

TRPC3 TRPM2 TRPP2 TRPA1 TRPML1 TRPV1

V D Y V QE S D - - - - S E V T - - - - - - - - - - - - - - - - - - T R NV EVL Q L D- - - T GI I F V L P - V - - - - S G - - - - I MK I G- I A - - - F V D Y S E M- F L QS L FM

PP - QN - KN AT

E T V

TRPC3 TRPM2 TRPP2 TRPA1 TRPML1 TRPV1

I AYI L L MH I - K I Q E C TF H RG QQM

TRPC3 TRPM2 TRPP2 TRPA1 TRPML1 TRPV1

V S

A A T

K K S

- - - - - - QA I L I HN - - - - - - - - - - - - - - - - - - H R WR GP A

TRPC3 TRPM2 TRPP2 TRPA1 TRPML1 TRPV1

P L K

E A R R A

S N N S -

DDA RV EL SL - -

T L A -

- HK- QQRP A GP I Y HP L S - - WL - - - -

TRPC3 TRPM2 TRPP2 TRPV1

E A

L I

S H S T

- D D I

I I D

- EE RTE

C M -

I V I -

A T A -

I L T -

WL F F - - Y

K Y -

CL CM SG - -

V L A -

F SY E E

NA FA FP - P

E GP D- P YVV - - - - RPV

L P FT - GI - GI

VL AF GY - -

S A L

V P Y -

V L L -

GL GL RT - -

- P I S RE - -

TT CL - - -

L F -

F F E -

- RE TA - -

P A -

NI T VT Y V L MV - I P S WQ - QDK W

I F I V

K A -

- L RL QV - -

QY - - PN P- - - L Y

A Q A -

- I DV SL - -

Y F F -

GY GKK - -

Q Q - -

K L -

WI - NV - -

A P T P W- -

CS AA - - -

RL R- L - -

GK I L - - - A DRGT - - - - -

Q K R

F I V

V V -

T V I

Q - T Y

I L F

R R K

K P Y R

V F ML S K A I L C - - - - V - M I - - - - - - N WY I I S L G- V

T Y ARDK - Y - - - - E A- Y A - - Q - A- SY S K- EY

L -

P P -

S G -

DP - QI QDV - V

T T F

T WT E S WC E DF F L NF L V

QI SE RV - - QF NNI A - - - - C CS L L A S M- - -

A E S F GP QI S G T V I K MV L MV F - MI I S K T L GP K I I V K MM V - - - - - - - - F L - L F R T MS Q S T T MS C A G AI M F I I L YAQ F- - - VM EV - - - - T L STVV F L L L GY I L AT RVA P- - - - SVM CCCV A V I Y L GY - CF YA VM EK M- - - - - R C MF V Y V F L F G S T A

Y K

H S

- - - V ERRV - - - - - - - - - CRPP

I P T A S I

- - GRP - - FL K

N D D

A A WL - - - S S

F F Y

T- T RGA - T- F - PL L M N L Y

E S YHS Q L SI S ST

ENI E WL T- Q- Q- - - -

GY TV - - - L - I

V YGI L L CL - FVF - VS Y YS L L A Y

Y Y I V

- AA - CM

P K

F F

- HL - L L

P R

P A

- I L - RS

L G

S K

F Y I I L L

K L F Q - EL

L I F

Q K

W GQ - R - M - L - T

NVMVV L L N- L MF - - - - I TI V - P SL I - YM - I L - Y L - QL - QV

F G

I Y

K T

P -

G -

F - Y D L - ML N - GM

L V -

SNF - - - - -

L

K D

D

Y

M I -

K D - -

ML CA - A Y C

I I A L

MV WV L G MMWS Y L WL S L V C E CEI I CFF- Y MI I T MA - -

G A A S C V

L L A A T V

Y V I S F

AFL - - I - - - - - - QRR

A I A V- S D - T V F V A V Y Y T - L V S A L G

A V A

HA - T YN YS S

QA PA - - - PS

T K - - - - MK

Y F Y V

SF PV NI SE

A -

I V N T

QQ - N T T L

- - V - - I I K- MN F V GV I T NM

SF - F L F L Y RY YY

S C L L T

MF L Y - - - - Y V WV - - - - - - Y L F GT Q V D F Y L L NL Q- P GW L GP Y HV K V T L I E DGK N L

YL - R P

G G E

- PK EE EP QQ - D

C Y G F

VE FA QI FQ Q- Q A E - - - - - - - - - - NI L Q

M R

- - EV T S V - - V L K NP E H C S P GT DP - - - - I A - - - - - I YR SF - D M F V T A A MQ - - - - L E T N

MI SS MF Y F L I I D M L - L - - - - - - - - - - - - - - - L MGE

- - - RVVL - - - P DGK

P T - -

I I AAF TA RF L A GL T C R I V L V A I GI - - - - N L E WI L L T D - - Y F F RGI QY F

V S -

L L W S S

QE QQ SE - - NK

I V V I

E Q K A

D E S Q

DS HT DL - - ES

D D A K

K I L A -

- - - - - - - - - R WC F

FIGURE 1.2 Amino acid sequence alignment of the ion channel domain of representative members of the TRP superfamily of cation channels: TRPC3, TRPM2, TRPP2, TRPA1, TRPML1, and TRPV1. Amino acids shown in white on a black background denote three identities among the six sequences. The consensus sequence is shown in bold. Note that there is signiﬁcant sequence similarity among members of the TRP channel superfamily.

(panels A through F of Figure 1.3) and located the positions of the exon boundaries. The ﬁgure panels show the deduced location of the putative transmembrane domains and channel pore-forming segment, and highlight exons coding for speciﬁc domains such as calmodulin binding, ankyrin repeats (absent in TRPM channels), IP3 receptor interacting sites, etc.

The TRPC Family of Ion Channels

9

FIGURE 1.3 Comparison of exon boundary locations along the open reading frames of TRP channels. A, TRPCs; B, TRPMs; C, TRPVs; D, TRPAs; E, TRPP; F, TRPML. Black boxes, 5′ and 3′ untranslated sequences; open boxes, coding exons; diagonally hatched rectangles, hydrophobic domains and pore regions; ovals, TRP motifs; hatched rectangles, hydrophobic domain. Adapted from Birnbaumer et al.35

The sequence alignments also revealed that C, M, and V TRP channels but not the TRPP, TRPML, or TRPA channels contain a typical six amino acid motif about ﬁfteen amino acids after the predicted sixth transmembrane segment of the TRP channel domain (Table 1.2), and that h1, the intracellular hydrophobic region that precedes the ion channel domain,52 is a feature found in all the TRP subfamilies with the exception of the TRPV subfamily. The cytosolic h domain also appears to be absent from some TRPs of the M, ML, and A1 classes. Domains and binding sites found in the various TRP channels are illustrated in Figure 1.4.

10

FIGURE 1.3 (CONTINUED).

TRP Ion Channel Function in Sensory Transduction

The TRPC Family of Ion Channels

11

FIGURE 1.3 (CONTINUED).

GENOMIC STRUCTURES Figure 1.5 illustrates the organization of the TRPC channels in the murine and human genomes, and that of the M-, V-, and P-type TRPs. The number of exons of each gene is given in Table 1.1. At this level of analysis, TRP genes show no special features. They extend from as little as 23.5 kb (mouse TRPC2) to as much as 304 kb (TRPC5). Large introns are spliced out both from within 5′ untranslated segments of the transcripts (e.g., intron A of TRPC5 = 164 kb) and from within the coding region of the transcripts (e.g., intron B of TRPC6 = 65.9 kb). The number of exons also varies from as many as over 40 (TRPP1) to as few as 2 (one coding and one 3′ noncoding: PKD-REJ [TRPP4]).

12

TRP Ion Channel Function in Sensory Transduction

TABLE 1.2 TRP Motifs TRPC Type

Sequence

TRPM Type

Sequence

TRPV Type

Sequence

C1 C2 C3 C4 C5 C6 C7

EWKFAR EWKFAR EWKFAR EWKFAR EWKFAR EWKFAR EWKFAR

M1 M2 M3 M4 M5 M6 M7 M8

VWKFQR IWKFQR VWKFQR YWKAQR FWKFQR LWKYNR VWKYQR VWKFQR

V1 V2 V3 V4 V5 V6

IWKLQR IWKLQR IWRLQR IWKLQW LWRAQV LWRAQI

TRPC Consensus EWKFAR

TRPM Consensus [VIYFL]WK[FAY][QN]R

TRPV Consensus [IL]W[KR][LA]Q[RWVI]

DIRECT CHROMOSOMAL REPEATS There may be two instances in which TRPs are found as direct repeats on a chromosome: TRPV3 (VRL2) follows TRPV1 (VR1), and TRPV6 (CaT1, ECaC2) follows after TRPV5 (ECaC1, CaT2), separated from the Stop codon of one and the beginning of the ORF of the next by only 7.5 Kb and 22 Kb, respectively. Both of these duplications appear to have been recent because, when compared to other

FIGURE 1.4 Overview of domains found in TRP channels. CaMBD, calmodulin-binding domains, are according to Zhang et al.,87 Tang et al.,88 Trost et al.,89 and Yildirim et al.90 IP3R Bdg Sites, inositol-trisphosphate receptor binding sites, are according to Boylay et al.65 Adapted from Birnbaumer et al.35

The TRPC Family of Ion Channels

13

FIGURE 1.5 Chromosomal organization of exons in genes coding for TRP channels. (Left) Murine and human TRPC genes. (Right) TRPM, TRPV, and TRPP genes. Exon predictions were obtained from genomic contigs downloaded from the National Center for Biotechnology Information (www.ncbi.nlm.nih.gov) human and murine genomic sequence databases. Exons coding for the ATG initiation codon of the open reading frame were arbitrarily numbered as 1 (one). Accession numbers and exact location of genes in each genome can be found in Table 1.1. Exons 3 and 11b of TRPC1 and TRPC4 are subject to alternative splicing and contain an ankyrin repeat and two CaM binding sites, respectively. Human TRPC2 is a pseudogene presenting “remnant” exons 2 and 3 ca. 70 Mb upstream of the main locus, missing exons 4 through 13, 15, and 16, but conserving the orthologous exons 14 and 17 through 21. (From Birnbaumer et al., Cell Calcium, 33, 419, 2003. With permission.)

TRPVs, V1 is most similar to V3 as compared to being most similar to another TRPV. The same applies to TRPV5 (ECaC1) and TRPV6 (CaT1). By comparison, in the ﬁeld of G protein α subunits, there are several pairs, such as Gi2α preceding Gtrα (locus 3p21), Gi3α preceding Gtcα (locus 1p13), Gi1α preceding Ggustα (locus 3p21), Gqα preceding G14α (locus 9p21), and G11α preceding G16α (locus 19p13). Giα’s are more similar to each other than to any of the transducins and gustducins, which in turn resemble each other much more than the Gi’s. Likewise Gqα is closer to G11α than to G14α, and G14α is closer to G16α than to G11α or Gqα, suggesting that gene duplications have happened very early in evolution, with duplicated genes duplicating further and allowing the leading and trailing member time to diverge.

14

TRP Ion Channel Function in Sensory Transduction

MECHANISM(S) OF TRPC ACTIVATION RECEPTOR-MEDIATED ACTIVATION OF TRPCS REQUIRES PHOSPHOLIPASE C ACTIVATION: A ROLE FOR PDZ SCAFFOLDS? The major players involved in regulation of TRPC channel activity are illustrated in Figure 1.6. A description of their involvement follows. TRPC channels are activated when phospholipase C is activated either by a Gq-coupled GPCR pathway mediated by PLCβs or by a receptor tyrosine kinase signaling pathway mediated by the γ family of PLCs. Activation of TRPCs is lost by inhibition of PLCβ with the PLCβ inhibitor U7312253 or when activation is tested in systems lacking PLCs, such

FIGURE 1.6 (Color ﬁgure follows p. 234.) Overview of some of the mechanisms that regulate or are thought to regulate the Gq-PLCβ triggered activation of TRPCs. A ligand of a Gq-coupled G-protein-coupled receptor (GPCR) is shown to activate the receptor’s Gqactivating function whereby Gq’s GDP is changed to GTP with concomitant dissociation of the heterotrimeric Gq protein into GTP-α plus the β-γ dimer. Both GTP-αq and β-γ independently activate β-type phospholipase Cs (PLCβs) leading to the hydrolysis of phosphatidylinositol bis-phosphate (PIP2) into inositol-trisphosphate (IP3) plus diacylglycerol (DAG). TRPC channels are depicted as being activated by three distinct mechanisms: (1) by DAG, presumably acting by direct interaction with the TRPC; (2) by the inositol trisphosphate receptor (IP3R), also thought to interact directly with the TRPC; and (3) possibly by STIM1, an ER Ca sensor that upon store depletion is translocated from the endoplasmic reticulum membrane (ER) to the plasma membrane (PM). The ﬁgure also highlights the negative regulation by PKC, activated by the cooperative interaction effect of DAG (generated by the action of the PLCβ) and Ca2+, originating ﬁrst from the store, and later from the extracellular milieu entering through the TRPC. CAMKs, Ca-calmodulin activated kinases; NOS, nitric oxide synthase; CN, calcineurin—also PP2B.

The TRPC Family of Ion Channels

15

as the NorpA Drosophila mutant, which lacks PLCβ,54 and DT40 chicken B cells, in which PLCγ has been inactivated by gene disruption.55 In Drosophila, the argument has been made for the formation of a ‘‘signalplex’’ with participation of INAD, a multi-PDZ domain–containing scaffold protein, as an intrinsic mechanism by which the trp/trpl-based phototransduction channel is activated. In agreement with this postulate, INAD binds to NorpA, PLC, trp, trpl, rhodopsin, and PKC56–59 (reviewed in reference 60). In mammals, including humans, two of the TRPCs—TRPC4 and TRPC5—also interact with a PDZ scaffold protein, NHERF, the regulator factor of the Na-H exchanger.61 NHERF is a two-PDZ domain protein. TRPC4, TRPC5, PLCβ1, and PLCβ2 interact with the ﬁrst PDZ domain, while the other PDZ domain binds members of the Ezrin-Radixin-Moesin (ERM) family of proteins, known to interact with F-actin. Thus, it would appear that, rather than organizing a signalplex similar to INAD’s role in the Drosophila eye, NHERF’s role in vertebrates may be that of physically connecting members of the PLC-TRP signaling pathway to the cytoskeleton. Yet this connection is unlikely to be part of the TRPC-activating process, because cortical actin, induced by calyculin A, blocks GPCR as well as storedepletion activated Ca2+ entry mediated by TRPC3.62 Calyculin A is a PP1 and PP2A phosphoprotein phosphatase inhibitor that causes accumulation of the C-terminally phosphorylated forms of ERM proteins. These in turn interact with F-actin, promoting its redistribution to the plasma membrane where the N-terminal portion of ERM proteins interact with membrane proteins. As a consequence, calyculin A treatment forms cortical actin. The implications of establishing independent connections between either PLCs and the actin cytoskeleton, or TRPC4-5 and the actin skeleton, which could interfere with TRPC’s function as an ion channel, are not clear. However, formation of transient complexes is likely to be involved in the process by which TRPCs are activated. Analysis of TRPC6 activation by Gq-coupled M1 muscarinic receptors in PC12D cells by Kim and Saffen63 showed the formation of time-sensitive macromolecular complexes involving PKC and phosphorylation of TRPC6 at a conserved PSPK site 23aa downstream of the TRPC EWFKAR motif. These authors found that once PKC was dissociated from the phosphorylated channel, the PS(PO3−)PK motif of TRPC6 recruits the FK506 binding protein FKBP12, which in turn recruits calcineurin (CN) and calmodulin (CaM). CN then dephosphorylates TRPC6, causing dissociation of the complex into its individual components. The complexes did not form if the Ser of the PSPK motif was mutated, when PKC was inhibited, when the immunophilin FKBP12 was blocked with FK506 or rapamycin, or when cells had been treated with cyclosporin. In all these instances, the channel was not dephosphorylated and the M1R stayed associated with the channel.

ACTIVATION BY CONFORMATIONAL COUPLING: THE ROLE FOR IP3 RECEPTOR AS AN ACTIVATOR OF TRPC THROUGH PROTEIN-PROTEIN INTERACTION The conformational coupling concept emerged originally from the elucidation of the role of the skeletal muscle voltage-gated Ca2+ channel in mediating a depolarizationinduced contraction. Skeletal muscle ﬁbers have a particularly strong Ca2+ uptake

16

TRP Ion Channel Function in Sensory Transduction

activity performed by SERCA pumps. As a consequence, the Ca2+ released from the sarcoplasmic reticulum (SR) in response to Ca2+-activated Ca2+ release through the ryanodine receptor/Ca2+ release channel is almost quantitatively reabsorbed into the SR with little or no loss through the action of plasma membrane Ca2+ pumps. Isolated skeletal muscle ﬁbers then contract repeatedly in response to repeated depolarizing stimuli. Initially, the Ca2+ that triggers Ca2+-activated Ca2+ release through the RYR was thought to come from the extracellular milieu, admitted through the skeletal muscle voltage-gated Ca2+ channel (CaV1.1). In support, depolarizing in the presence of a CaV1.1 channel blocker, such as a dihydropyridine (DHP), abolished the response to depolarization. It was surprising that contractions that were blocked by DHPs could still be elicited in totally Ca2+-free media. Activation of the RYR did not depend on an initial Ca2+ inﬂux through the channel but did nevertheless depend on the presence of a fully functional voltage and DHP-sensitive complex. The molecular basis for this phenomenon became clear when it was discovered that upon membrane depolarization, the Ca2+ channel changed its conformation and interacted physically with the nearby RYR, triggering its initial opening, initial Ca2+ release, and the ensuing explosive Ca2+-activated Ca2+ release responsible for activation of the actomyosin contractile machinery (reviewed in reference 64). The conformational coupling model for TRPC activation is based on the skeletal muscle excitation-coupling model, in which the information ﬂow is from a plasma membrane ion channel (CaV1.1) to a endomembrane Ca2+ release channel (RYR), but with the ﬂow of information (i.e., the sense of the signaling pathway) inverted. The model postulates that IP3, the same stimulus that activates the IP3R to release Ca2+ from the endoplasmic reticulum Ca2+ stores, also activates a TRPC-activating function of the IP3 receptor. Activation of TRPCs would come about by physical binding of a TRPC-binding domain of the IP3R to TRPC with attendant activation of the affected TRPC. This hypothesis was tested in our laboratory in the late 1990s using a GST pulldown approach.65 We found that a region on the post-transmembrane C-terminal domain of TRPCs interacts with a region located between (IP3R-3 numbering) amino acids 675 and 800 of the IP3Rs. This lies about 100aa C-terminal to the IP3 binding domain (aa 225–575) distal to the C-terminally located ion channel forming transmembrane domains (aa 224–2565) of the 2761 aa IP3R-2. More important, transient overexpression of GST-fusion fragments of TRPC-interacting sequences of the IP3R either inhibited or extended the duration of Ca2+ inﬂux through endogenous receptoror store depletion–activated Ca2+ entry channels.65 Thus, IP3R sequences identiﬁed as TRPC binding sequences affect Ca2+ entry that is postulated to be mediated by TRPC channels. The regions identiﬁed on IP3R-2 are conserved in IP3R-1 and IP3R-3, and the sequences identiﬁed on TRPC3 as interacting with IP3Rs are preserved in other TRPCs. Researchers in Sage’s laboratory66 used a different approach to test the conformational coupling hypothesis. Working with human platelets, they probed for co-immunoprecipitation of endogenous TRPC1 with endogenous IP3R. No IP3R co-immunoprecipitated from control lysates of control platelets, but activation of Ca2+ inﬂux secondary to store depletion, induced by inactivation of SERCA pumps with thapsigargin, resulted in IP3R co-immunoprecipitating with the human TRPC1.

The TRPC Family of Ion Channels

17

As was the case in earlier experiments with A7r5 and DDT1-MF2 smooth muscle cell lines,62 induction of cortical actin in platelets with jasplakinolide blocked the interaction of the IP3R with TRPC1. These experiments are especially relevant because the activation-dependent interaction of IP3R with TRPC was shown in a normal cell with normal complements of the interacting partners. Interactions observed under these conditions do not suffer from the drawbacks of interactions shown only on overexpression of the interacting partners. It remains to be determined how IP3Rmediated activation of TRPC channels interfaces with STIM1 translocation and activation of Icrac.

ACTIVATION OF TRPCS BY DIACYLGLYCEROL (DAG) TRPCS BY PROTEIN KINASE C (PKC)

AND INHIBITION

OF

In 1999, Hofmann et al.67 reported that diacylglycerols, one of the two reaction products resulting from PLC activation, activate TRPC3 and TRPC6, and that they do so independently of PKC activation. That same year, Okada et al.25 reported the same ﬁnding for the newly cloned TRPC7. In both studies the effects of DAGs could be augmented by inhibiting DAG lipase or DAG kinase, thus sparing DAG removal, and were insensitive to PKC inhibitors. The study by Okada et al.25 also showed that activation of PKC inhibits activation of TRPC7 by subsequently adding DAG and indicating opposite roles for PKC and DAG. A superﬁcial examination of DAG’s effects on TRPC4 and TRPC5 does not show stimulation by DAG. Yet upon inhibition of PKCs, both TRPCs are robustly activated by DAG.25,55 Indeed, activation of PKC inactivates all TRPCs so far studied for this effect (TRPC3 through TRPC7), not only for activation by DAG but also by the Gq-coupled and the RTK (receptor tyrosine kinase) signaling pathways.25,55,68 But, the rates at which DAG and PKC act on the TRPCs differ with TRPC subtype. Thus, for the TRPC3 family of TRPCs (TRPC3, TRPC6, and TRPC7), activation by DAG is faster than phosphorylation by PKC so that activation by DAG is the dominant phenomenon. On the other hand, for TRPC4 and TRPC5, the DAG-induced inhibition by PKC is established before DAG can activate the TRPC channel. It has not been reported whether this differs after induction of cortical actin. DAG also activates the TRPC2 channel in its native environment, the vomeronasal sensory neuron.69 The effect is fast, potentiated by DAG lipase and DAG kinase inhibitors and unaffected by PKC inhibitors. Whether PKC is inhibitory to TRPC2, as it is for TRPC3 through TRPC7, has not been reported. Given that the Ser phosphorylated by PKC is located in a motif that is conserved in all TRPCs,68 it is tempting to suggest that PKC is inhibitory to all. In agreement with the interpretation that the channel being activated by DAG in vomeronasal sensory neurons is indeed TRPC2, genetic ablation of TRPC2 resulted in loss of the DAG-activated current.69

ACTIVATION OF CA2+ ENTRY BY CHANNEL TRANSLOCATION FROM ENDOMEMBRANES TO THE PLASMA MEMBRANE Insulin stimulates glucose uptake into its target tissues by promoting the translocation of GLUT4-bearing endovesicles to the plasma membrane. The incorporation

18

TRP Ion Channel Function in Sensory Transduction

of AMPA-type glutamate receptors into the postsynaptic membrane of neurons undergoing high-frequency stimulation is the basis for the establishment of early long-term memory (eLTP) in hippocampal CA1 neurons. The TRPV2 channel, also called the growth factor–regulated channel or GRC, transitions from internal membranes to the plasma membrane after myotubes are treated with insulin-like growth factor-1 (IGF-1). Boulay and coworkers70 tested whether TRPC6 might be under similar control and indeed saw an increase of cell surface TRPC6, as seen by an increase of biotinylated TRPC6 by immunostaining within 30 sec of stimulating cells either with an agonist for a Gq-coupled GPCR (M5R) or with the store-depleting agent thapsigargin. In a different context, Clapham and collaborators showed that activation of Rac1 in hippocampal neurons leads to translocation of TRPC5 from endomembranes to the plasma membrane in a process that involves activation by Rac1 of PIP(5)K (phophatidylinositol-4-phosphate 5-kinase) and synthesis of PIP2 (phosphatidylintositol-4,5-bisphosphate). Rac1 in turn is activated by stimulation of an RTK (e.g., EGFR) activating PI3K with formation of PIP3 (phosphatidyl-inositol-3,4,5-triphosphate), which in turn activates a Rac1 guanine nucleotide exchange factor (Rac-GEF), leading to augmented GTP-Rac1.71 This signaling mechanism (RTK to TRPC5 incorporation into plasma membrane) has been implicated in Ca2+-dependent repression of neurite outgrowth, because there is an inverse correlation of PIP(5)K with neurite length.

PHYSIOLOGICAL ROLE FOR TRPC CHANNELS AS ELECTROGENIC DEVICES THAT COUPLE GPCR-GQ ACTIVATION TO VOLTAGE-GATED CA2+ CHANNEL ACTIVATION Activation of nonselective monovalent cation channels leads to a collapse of the membrane potential. Many TRP and TRP-related channels are mostly nonselective cation channels, some with selectivity for monovalent cations over divalents, some nonselective with respect to both monovalent and divalent cations. As such, activation of these types of channels dissipates the transmembrane potential of the cells in which the channels are expressed. This property was highlighted in a report from the Fleig-Penner laboratory in which the channel properties of TRPM5, a Ca2+activated nonselective monovalent cation channel (i.e., a CAN), was characterized.72 More recently, Soboloff et al.,73 studying the effect of down-regulating TRPC6 with small interfering RNA (siRNA), found that, in A7r5 smooth muscle cells, TRPC6 fulﬁlls the role of an electrogenic coupling mechanism by coupling a Gq-coupled GPCR to Ca2+ inﬂux through a dihydropyridine-sensitive Ca2+ channel. This became apparent when, upon siRNA treatment, they saw a >90 percent reduction of muscarinic receptor-stimulated TRPC6 channel activity, measured by the patch clamp technique, with essentially no loss of Ca2+ inﬂux, measured with the ﬂuorescent Ca2+ indicator dye, fura2. Muscarinic receptor-stimulated Ca2+ inﬂux into siRNA-treated cells was completely inhibited by a dihydropyridine Ca2+ channel blocker (see reference 73). Although this is the ﬁrst demonstration of an electrogenic coupling role for a TRPC channel, it is likely that other examples are soon to follow, especially in natural tissue cells.

The TRPC Family of Ion Channels

19

A ROLE FOR TYROSINE KINASES IN VOLTAGE-GATED CA2+ CHANNEL-INDEPENDENT CA2+ INFLUX The original observation that tyrosine phosphorylation may be an important regulator participating in activation of receptor- and store-operated Ca2+ entries (ROCE and SOCE) in nonexcitable cells came from studies showing an inhibitory effect of tyrosine kinase inhibitors on these forms of calcium entry in human foreskin ﬁbroblasts.74,75 This was followed by studies that showed the total loss of bradykinin-stimulated ROCE and partial absence of SOCE in embryonic ﬁbroblasts derived from mice lacking the src tyrosine kinase.76 Given that evidence has accumulated showing that tyrosine phosphorylation is a common consequence associated with cell stimulation via receptors that signal by using the Gq-PLCcalcium mobilizing pathway,77,78 we became interested in the possibility that tyrosine phosphorylation may be an activating signal for one of the events that leads to receptor- or store-operated calcium entry. Indeed, experiments parallel to ours that tested for a role of tyrosine kinases in the receptor-mediated activation of the type 3 TRPC (TRPC3) revealed that the activation of this TRPC by a PLCstimulating GPCR in HEK cells is inhibited by inhibitors of tyrosine kinases; when expressed in src kinase–negative cells, the transfected TRPC3 is not activated by a cotransfected Gq-coupled receptor.79 This recapitulated the earlier ﬁndings with an endogenous (bradykinin-activated) GPCR acting via Gq activation on the endogenous complement of the receptor-operated Ca2+ entry pathway.76

STUDIES ON THE ROLE(S) OF TYROSINE PHOSPHORYLATION IN TRPC FUNCTION During the last few years and in order to learn more about the possible role of tyrosine kinase(s) in TRPC-mediated events, as well as in SOCE, we used in vitro and in cell protein–protein interaction assays.80 We found that c-src phosphorylates TRPC3 on tyrosine 226 (Y226) located on the TRPC3 N-terminus and that formation of phospho-Y226 is essential for TRPC3 activation. Surprisingly, this requirement is unique for TRPC3, because (1) mutation of the cognate tyrosines of the closely related TRPC6 and TRPC7 channels had no effect on their function; (2) both TRPC6 and TRPC7 were activated in src-, yes-, and fyn-deﬁcient cells; and (3) src, but not yes or fyn, rescued TRPC3 activation in cells lacking src, yes, and fyn. Yet we found the SH2 domain of c-src to interact not only with TRPC3 but also with either the N-termini or the C-termini of all other TRPCs. This suggests that other tyrosine kinases may play a role in ion ﬂuxes mediated by TRPCs. A side-by-side comparison of the effects of genistein on endogenous ROCE and SOCE in YF, SYF, HEK, and COS-7 cells showed these inﬂuxes to be inhibitable in all three cell types but with differing sensitivities (Figures 1.7 and 1.8). Taken together, these results argue for the channels mediating ROCE and SOCE to be heterogeneous and to differ from tissue to tissue. The ﬁnding that TRPC6 is active in SYF and YF cells was unexpected, as it has been shown to be a substrate of fyn81 and to behave essentially the same as TRPC3

20

TRP Ion Channel Function in Sensory Transduction

FIGURE 1.7 Comparison of the inhibitory effect of genistein on ROCE and SOCE endogenous to HEK cells, COS-7 cells, and mouse embryonic ﬁbroblasts lacking the indicated members of the src-family of tyrosine kinases. In the experiments shown in this and the other ﬁgures, ROCE was assessed in cells expressing the indicated Gq-coupled GPCR in transient or stable form, loading the cells that had been plated on coverslips with fura2 and subjecting these cells to a Ca2+ mobilization protocol in which the PLC system was activated by the cognate receptor agonist (carbachol [CCh] for the M5 muscarinic receptor and arginine vasopressin [AVP] for the V1a vasopressin receptor) in the absence of external Ca2+. [Ca2+]i changes were then followed by video spectromicroscopy to record Ca2+ release from internal stores and allowing [Ca2+]i to return to near basal levels. At this point, Ca2+ was added to the external medium and inﬂux of Ca2+ leading to increase in [Ca2+]i representing ROCE, was monitored for the indicated times. When drugs such as genistein or KB-R7943 (see below) were added, they were present through the ﬁrst and second phases of the [Ca2+]i changes. Note that only the Ca2+ entry phases are shown. In none of the experiments shown in this or the other ﬁgures did the presence of tyrosine kinase or Na-Ca exchange inhibitors affect signiﬁcantly the IP3- or thapsigargin-induced Ca2+ release from the endogenous stores. SOCE was assessed by substituting the GPCR agonist for thapsigargin. Gd3+ (5–10 μM) was added when TRPC-mediated ROCE was measured. At this concentration of Gd3+ TRPC3, 5, 6, and 7 mediated Ca2+ entry is unaffected, but endogenous ROCE is inhibited. Data on effects of genistein are either unpublished or adapted from Kawasaki et al.80

in in vitro and cell expression assays. Thus, TRPC6 is phosphorylated by coexpression with fyn in COS cells, and it associates with fyn in GST pull-down assays by interaction of its N-terminus with the SH2 domain of fyn,81 and TRPC6 activation is inhibited by the tyrosine kinase inhibitor PP2, regardless whether it is activated by a receptor-tyrosine kinase-PLCγ pathway (triggered by EGF)81 or by the DAG pathway.73 Further, addition of fyn to inside-out membrane patches from cells expressing TRPC6 increased basal and oleyl-acetyl-glyceride (OAG)–stimulated

The TRPC Family of Ion Channels

21

FIGURE 1.8 Comparison of the inhibitory effect or lack thereof of genistein on ROCE mediated by the indicated TRPC channels as seen in HEK cells. Adapted from Kawasaki et al.80

TRPC6 activity.81 Yet, TRPC6 ROCE is activated in cells lacking not only fyn but also yes and src (Figures 1.7 and 1.8). Not known at this time is whether this discrepancy is either due to our use of the GPCR-Gq-PLCβ activation pathway instead of a RTK-PLCγ pathway (which might be impaired in SYF cells) or due to the different form of assessing TRPC6 ROCE (fura2 in our case and electrophysiological in inside-out membrane patches in the case of Hisatsune et al.).81 If, however, the lack of sensitivity of TRPC6 to genistein in our experiments and the activity of TRPC6 in SYF cells is not due to the use of differing activation pathways or to the method used to assess TRPC6 activation, the data may also indicate that functionally, in addition to src, fyn, and yes, there is at least one more “PP2-sensitive src-family tyrosine kinase” able to regulate TRPC6 and other TRPCs. As mentioned, the fact that TRPC1 through C7 all interacted with src in GST pull-down assays raises the possibility that all TRPC channels depend on tyrosine phosphorylation for their functioning as an effector system for the activation of the GPCR-Gq-PLCβ/RTKPLCγ pathways. Table 1.3 summarizes properties of TRPC channels in terms of which signaling pathways have been shown to activate each channel and also possible mechanisms by which each is activated. Two pathways feed into TRPCs. One generates DAG from the action of PLCβ-activated by Gq and Gi-derived Gβγ, and the other generates DAG from the action of PLCγ-activated by tyrosine phosphorylation either directly by the RTK-type receptor or secondarily to non-RTK tyrosine kinase recruitment such as happens with T- and B-cell receptor activation. Activation of TRPCs by DAG may be aided by IP3R acting by protein–protein interaction according to a conformational coupling model and by the formation of a multimolecular signaling complex with or without involvement of a protein acting as a nucleating scaffold. A direct IP3R–TRPC interaction, especially if facilitated by the action STIM1,10–12 may also account for activation of TRPC channels by store depletion.

yes

yes97

? yes***25,98,99

TRPC1 TRPC2 TRPC3 TRPC3a TRPC4

TRPC5

TRPC6 TRPC7

Inh.55 Inh.73 Inh.25,98,99

DAG67 DAG25,98,99 yes yes

yes

yes yes yes yes yes

Activation by RTK#

Inhibition Inhibition

no80, yes73 no80

no80

? ? yes-Y22680 nt ?

Inhibition by Genistein PP2

no yes

no no

no

no no no no no

Highly Ca2+ Selective?

EGF receptor (EGF), B cell receptor (anti-IgM), T cell receptor (anti-CD3). * thapsigargin; ** M5 muscarinic or V1a vasopressin receptors; *** requires low level of expression. Inh., inhibition; Stimul’n, stimulation; Conformat’l, conformational; nt, not tested; HSWP, human foreskin ﬁbroblasts.

#

? ? Inh.55 nt Inh.55

Effect of PKC

DAG55

? DAG69 DAG67 nt DAG55

2nd Messenger

ROCE (Gq-coupled Rs): HEK, YF MEF, COS, HSWP74 SOCE (thapsigargin): HEK, YF MEF, COS, HSWP

yes yes

yes yes yes yes yes

yes66,91 yes92,93 yes***94 yes***95 yes96

Channel

Stimul’n by GqCoupled GPCR**

Activation by Store Depletion*

Inhibition Inhibition

Inh.? Inh.? Resistant Resistant ? Activated @100 μM100 Resistant Activated @100 μM101 Resistant,80 Inh.102 Resistant

Channel Blocked by Gd3+ (5-10 μM)

yes yes

yes

yes yes yes yes yes

Conformat’l Coupling (Interaction with IP3Rs)

TABLE 1.3 Regulation of TRPC Channels: Comparison to Endogenous ROCE and SOCE of HEK Cells, yes- and fyn-Deﬁcient Mouse Embryonic Fibroblasts (YF MEFs), COS-7 Cells, and HSWP Human Foreskin Fibroblasts

22 TRP Ion Channel Function in Sensory Transduction

The TRPC Family of Ion Channels

TRPCS

AND

23

SOCE

In contrast to the rather satisfactory models one can set up for explaining how TRPC channels may be activated (Figure 1.6), there are only scant data that suggest how, if at all, TRPC channels participate in SOCE. The strongest data set was recently published by Villereal studying the natural channels that contribute to SOCE in HEK293 cells testing for interference with siRNA. More than 80 percent downregulation of the naturally expressed TRPC1, TRPC3, and TRPC7 proteins resulted in a ca. 50 percent reduction in thapsigargin-induced SOCE.82 The 50 percent loss is curious, because it happens to coincide with the similar loss of acetyl choline–induced NO-mediated vascular relaxation of aortic rings seen in mice lacking TRPC4.83 The missing information or hypothesis relating TRPCs to SOCE is how nonselective cation channels may come together to form a highly Ca2+-selective ion channel. On a purely speculative level there are three possibilities that come to mind. One is that ion selectivity does change in channels that are heteromeric in nature. The other is that ion selectivity is altered by post-translational modiﬁcation. Dietrich et al.84 have shown that basal or constitutive activity is affected by glycosylation. Kawasaki et al.80 found that at least one channel, TRPC3, depends on tyrosine phosphorylation for activity. The effect of compound kinase actions has not been explored. Groschner and collaborators, by showing that in HEK cells TRPC3mediated Ca2+ inﬂux depends on extracellular Ca2+ and a functional Na-Ca exchanger operating in reverse,85 raised the possibility that the Ca2+ selectivity is the result of a tandem arrangement whereby Na+ entering through a TRPC channel is extruded not only by the Na-K ATPase but also by the Na-Ca exchanger, an exquisitely Ca2+selective machine. The third possibility regarding the molecular makeup of ROCE, and especially SOCE channels, is that the ﬁeld may have been somewhat naive in assuming that there is only one Icrac channel. The fact that inhibition of endogenous SOCE by genistein shows varying degrees of sensitivity may indicate that SOCE channels— presumed to be equivalent to Icrac channels—are heterogenous in nature. If so, TRPrelated channels other than TRPCs may form SOCE channels. Participation of various TRPVs, especially TRPV5 and TRPV6, comes to mind. In line with this reasoning, Schindl et al.86 noted commonalities between TRPV6 (CaT1) and Icrac. The studies reported by Rosker et al.,85 implicating the Na-Ca exchanger in ROCE mediated by TRPC3 in HEK cells, prompted us to test the effectiveness with which the Na-Ca exchanger (NCX) inhibitor KB-R7943 affects ROCEs mediated by several TRPCs in HEK cells and ROCEs mediated by endogenous ROCE channels in HEK and other cells. The picture that emerged is not one to support an obligatory role for Na-Ca exchangers in ROCE or SOCE—for this KB-R7943 is too nonspeciﬁc— but one that suggests that endogenous channels mediating ROCE and SOCE are likely to be heterogeneous in their molecular makeup (Figure 1.9). ROCE and SOCE are forms of Ca2+ entry with speciﬁc functions in a large list of diverse cellular functions that include smooth muscle contraction, B- and T-cell activation, and vascular permeability and development of the central nervous system, to name a few. Yet, the molecular makeup of the channels mediating ROCE and

24

TRP Ion Channel Function in Sensory Transduction

FIGURE 1.9 Comparison of the effect of the partially selective Na-Ca exchange inhibitor, KB-R7943, on ROCE mediated by TRPCs and on ROCE and SOCE mediated by endogenous channels.

SOCE is still largely a matter of speculation. New tools are needed to conﬁrm or negate the hypothesis that TRPCs alone or in combination with other members of the TRP superfamily participate in these forms of Ca2+ entry.

ACKNOWLEDGMENTS Research was supported by the Intramural Research Program of the National Institutes of Health and the National Institute of Environmental Health Sciences.

REFERENCES 1. Pak, W.L., Grossﬁeld, J., and Arnold, K., Mutant of the visual pathway of Drosophila melanogaster, Nature, 227, 518, 1970. 2. Hotta, Y. and Benzer, S., Genetic dissection of the Drosophila nervous system by means of mosaics, Proc. Natl. Acad. Sci. USA, 67, 1156, 1970. 3. Montell, C. and Rubin, G.M., Molecular characterization of the Drosophila trp locus: a putative integral membrane protein required for phototransduction, Neuron, 2, 1313, 1989. 4. Phillips, A.M., Bull, A., and Kelly, L.E., Identiﬁcation of a Drosophila gene encoding a calmodulin-binding protein with homology to the trp phototransduction gene, Neuron, 8, 631, 1992. 5. Hu, Y., Vaca, L., Zhu, X., Birnbaumer, L., Kunze, D., and Schilling, W.P., Appearance of a novel Ca2+-inﬂux pathway in Sf9 insect cells following expression of the transient receptor potential-like (trpl) protein of Drosophila, Biochem. Biophys. Res. Commun., 132, 346, 1994.

The TRPC Family of Ion Channels

25

6. Devary, O., Heichal, O., Blumenfeld, A., Cassel, D., Suss, E., Barash, S., Rubinstein, C.T., Minke, B., and Selinger, Z., Coupling of photoexcited rhodopsin to inositol phospholipid hydrolysis in ﬂy photoreceptors, Proc. Natl. Acad. Sci. USA, 84, 6939, 1987. 7. Selinger, Z. and Minke, B., Inositol lipid cascade of vision studied in mutant ﬂies, Cold Spring Harbor Symp. Quant. Biol., 53, 333, 1988. 8. Hu, Y. and Schilling, W.P., Receptor-mediated activation of recombinant trpl expressed in Sf9 insect cells, Biochem. J., 305, 605, 1995. 9. Birnbaumer, L., Zhu, X., Jiang, M., Boulay, G., Peyton, M., Vannier, B., Brown, D., Platano, D., Sadeghi, H., Stefani, E., and Birnbaumer, M., On the molecular basis and regulation of cellular capacitative calcium entry: roles for Trp proteins, Proc. Natl. Acad. Sci. USA, 93, 15195, 1996. 10. Liou, L., Kim, M.L., Jones, J.T., Myers, J.W., Ferrel, Jr., J.E., and Meyer, T., STIM is a Ca2+ sensor essential for Ca2+-store-depletion-triggered Ca2+ inﬂux, Curr. Biol., 15, 1235, 2005. 11. Roos, J., DiGregorio, P.J., Yeromin, A.V., Ohlsen, K., Lioudyno, M., Zhang, S., Safrina, O., Kozak, J.A., Wagner, S.L., Cahalan, M.D., Vilicelebi, G., and Stauderman, K.A., STIM1, an essential and conserved component of store-operated Ca2+ channel function, J. Cell Biol., 169, 435, 2005. 12. Zhang, S.L, Yu, Y., Roos, J., Kozak, J.A., Deerinck, T.J., Ellisman, M.H, Stauderman, K.A., and Cahalan, M.D., STIM1 is a Ca2+ sensor that activates CRAC channels and migrates from the Ca2+ store to the plasma membrane, Nature, 437, 902, 2005. 13. Kwan, C.Y., Takemura, H., Obie, J.F., Thastrup, O., and Putney, J.W. Jr., Effects of MeCh, thapsigargin, and La3+ on plasmalemmal and intracellular Ca2+ transport in lacrimal acinar cells, Am. J. Physiol., 258, C1006, 1990. 14. Thastrup, O., Cullen, P.J., Drobak, B.K., Hanley, M.R., and Dawson, A.P., Thapsigargin, a tumor promoter, discharges intracellular Ca2+ stores by speciﬁc inhibition of the endoplasmic reticulum Ca2+ ATPase, Proc. Natl. Acad. Sci. USA, 87, 2466, 1990. 15. Hoth, M. and Penner, R., Depletion of intracellular calcium stores activates a calcium current in mast cells, Nature, 355, 353, 1992. 16. Putney, J.W., Jr., A model for receptor-regulated calcium entry, Cell Calcium, 7, 1, 1986. 17. Putney, J.W., Jr., Capacitative calcium entry revisited, Cell Calcium, 11, 611, 1990. 18. Zweifach, A. and Lewis, R.S., Mitogen-regulated Ca2+ current of T lymphocytes is activated by depletion of intracellular Ca2+ stores, Proc. Natl. Acad. Sci. USA, 90, 6295, 1993. 19. Hardie, R.C. and Minke, B., The trp gene is essential for a light-activated Ca2+ channel in Drosophila photoreceptor cells, Neuron, 8, 643, 1992. 20. Hardie, R.C. and Minke, B., Novel Ca2+ channels underlying transduction in Drosophila photoreceptors: implications for phosphoinositide-mediated Ca2+ mobilization, Trends in Neurosci., 9, 371, 1993. 21. Sinkins, W.G., Vaca, L., Hu, Y., Kunze, D.L., and Schilling, W.P., The COOH-terminal domain of Drosophila TRP channels confers thapsigargin sensitivity, J. Biol. Chem., 271, 2955, 1996. 22. Zhu, X., Chu, P.B., Peyton, M., and Birnbaumer, L., Molecular cloning of a widely expressed human homologue for the Drosophila trp gene, FEBS Lett., 373, 193, 1995. 23. Wes, P.D., Chevesich, J., Jeromin, A., Rosenberg, C., Stetten, G., and Montell, C., TRPC1, a human homolog of a Drosophila store-operated channel, Proc. Natl. Acad. Sci. USA, 92, 9652, 1995.

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TRP Ion Channel Function in Sensory Transduction 24. Zhu, X., Jiang, M., Peyton, M.J., Boulay, G., Hurst, R., Stefani, E., and Birnbaumer, L., trp, a novel mammalian gene family essential for agonist-activated capacitative Ca2+ inﬂux, Cell, 85, 661, 1996. 25. Okada, T., Inoue, R., Yamazaki, K., Maeda, A., Kurosaki, T., Yamakuni, T., Tanaka, I., Shimizu, S., Ikenaka, K., Imoto, K., and Mori, Y., Molecular and functional characterization of a novel mouse TRP homologue TRP7 that forms a background and receptor-activated Ca2+ permeable cation channel, J. Biol. Chem., 274, 27359, 1999. 26. Montell, C., Birnbaumer, L., Flockerzi, V., Bindels, R.J., Caterina, M.J., Clapham, D.E., Heller, S., Julius, D., Scharenberg, A.M., Schultz. G., and Zhu, M.X., A uniﬁed nomenclature for the superfamily of TRP cation channels, Mol. Cell, 9, 229, 2002. 27. Hofmann, T., Schaefer, M., Schultz, G., and Gudermann, T., Subunit composition of mammalian transient receptor potential channels in living cells, Proc. Natl. Acad. Sci. USA, 99, 7461, 2002. 28. Lintschinger, B., Balzer-Geldsetzer, M., Baskaran, T., Graier, W.F., Romanin, C., Zhu, M.X., and Groschner, K., Coassembly of trp1 and trp3 proteins generates diacylglycerol- and Ca2+-sensitive cation channels, J. Biol. Chem., 275, 27799, 2000. 29. Struebing, C., Krapivinsky, G., Krapivinsky, L., and Clapham, D.E., Formation of novel TRPC channels by complex subunit interactions in embryonic brain, J. Biol. Chem., 278, 39014, 2003. 30. Caterina, M.J., Schumacher, M.A., Tominaga, M., Rosen, T.A., Levine, J.D., and Julius, D., The capsaicin receptor: a heat-activated ion channel in the pain pathway, Nature, 389, 816, 1997. 31. Caterina, M.J., Rosen, T.A., Tominaga, M., Brake, A.J., and Julius, D., A capsaicinreceptor homologue with a high threshold for noxious heat, Nature, 398, 436, 1999. 32. Kanzaki, M., Zhang, Y.-Q., Mashima, H., Li, L., Shibata, H., and Kojima, I., Translocation of a calcium-permeable cation channel induced by insulin-like growth factorI, Nature Cell Biol., 1, 165, 1999. 33. Hunter, J.J., Shao, J., Smutko, J.S., Dussault, B.J., Nagle, D.L., Woolf, E.A., Holmgren, L.M., Moore, K.J., and Shyjan, A.W., Chromosomal localization and genomic characterization of the melastatin gene (Mlsn 1), Genomics, 54, 116, 1998. 34. Veldhuisen, B., Spruit, L., Dauwerse, H.G., Breuning, M.H., and Peters, D.J.M., Genes homologous to the autosomal dominant polycystic kidney disease genes (PKD1 and PKD2), Eur. J. Hum. Genet., 7, 860, 1999. 35. Birnbaumer, L., Yildirim, E., and Abramowitz, J., A comparison of the genes coding for canonical TRP channels and their M, V, and D relatives, Cell Calcium, 33, 419, 2003. 36. Sun, M., Goldin, E., Stahl, S., Falardeau, J.L., Kennedy, J.C., Acierno, J.S., Bove, C., Kaneski, C.R., Nagel, J., Bromley, M.C., Colman, M., Schiffmann, R., and Slaugenhaupt, S.A., Mucolipidosis type 4 is caused by mutations in a gene encoding a novel transient receptor potential channel, Hum. Mol. Genet., 9, 2471, 2000. 37. Story, G.M., Peier, A.M., Reeve, A.J., Eid, S.R., Mosbacher, J., Hricik, T.R., Earley, T.J., Hergarden, H.C., Andersson, D.A., Hwang, S.W., McIntyre, P., Jegla, T., Bevan, S., and Patapoutian, A., ANKTM1, a TRP-like channel expressed in nociceptive neurons, is activated by cold temperatures, Cell, 112, 819, 2003. 38. Corey, D.P., Garcia-Anoveros, J., Holt, J.R., Kwan, K.Y., Lin, S.-Y., Vollrath, M.A., Amalﬁtano, A., Cheung, E.L.-M., Derﬂer, B.H., Duggan, A., Geleoc, G.S.G., Gray, P.A., Hoffman, M.P., Rehm, H.L., Tamasauskas, D., and Zhang, D.S., TRPA1 is a candidate for the mechanosensitive transduction channel of vertebrate hair cells, Nature, 342, 723, 2004.

The TRPC Family of Ion Channels

27

39. Hoenderop, J.G., van der Kemp, A.W., Hartog, A., van de Graaf, S.F., van Os, C.H., Willems, P.H., and Bindels, R.J., Molecular identiﬁcation of the apical Ca2+ channel in 1,25-dihydroxyvitamin D3-responsive epithelia, J. Biol. Chem., 274, 8375, 1999. 40. Peng, J.B., Chen, X.Z., Berger, U.V., Vassilev, P.M., Tsukaguchi, H., Brown, E.M., and Hediger, M.A., Molecular cloning and characterization of a channel-like transporter mediating intestinal calcium absorption, J. Biol. Chem., 274, 22739, 1999. 41. Yue, L., Peng, J.B., Hediger, M.A., and Clapham, D.E., CaT1 manifests the pore properties of the calcium release-activated calcium channel, Nature, 410, 705, 2001. 42. Fleig, A. and Penner, R., The TRPM ion channel subfamily: molecular, biophysical and functional features, Trends Pharmacol. Sci., 25, 633, 2004. 43. McKemy, D.D., Neuhausser, W.M., and Julius, D., Identiﬁcation of a cold receptor reveals a general role for TRP channels in thermosensation, Nature, 416, 52, 2002. 44. Qiu, X., Kumbalasiri, T., Carlson, S.M., Wong, K.Y., Krishna, V., Provencio, I., and Berson, D.M., Induction of photosensitivity by heterologous expression of melanopsin, Nature, 433, 745, 2005. 45. Panda, S., Nayak, S.K., Campo, B., Walker, J.R., Hogenesch, J.B., and Jegla, T., Illumination of the melanopsin signaling pathway, Science, 307, 600, 2005. 46. Wang, G.X. and Poo, M.M., Requirement of TRPC channels in netrin-1-induced chemotropic turning of nerve growth cones, Nature, 434, 898, 2005. 47. Li, Y., Jia, Y.C., Cui, K., Li, N., Zheng, Z.Y., Wang, Y.Z., and Yuan, X.B., Essential role of TRPC channels in the guidance of nerve growth cones by brain-derived neurotrophic factor, Nature, 434, 894, 2005. 48. Liman, E.R., Corey, D.P., and Dulac, C., TRP2, a candidate transduction channel for mammalian pheromone sensory signaling, Proc. Natl. Acad. Sci. USA, 96, 5791, 1999. 49. Stowers, L., Holy, T.E., Meister, M., Dulac, C., and Koentges, G., Loss of sex discrimination and male-male aggression in mice deﬁcient for TRPC2, Science, 295, 1493, 2002. 50. Dulac, C. and Torello, A.T., Molecular detection of pheromone signals in mammals: from genes to behavior, Nature Rev. Neurosci., 4, 551, 2003. 51. Zufall, F., Ukhanov, K., Lucas, P., and Leinders-Zufall, T., TRPC2: from gene to behavior, Pﬂuegers Arch. Eur. J. Physiol., 451, 61, 2005. 52. Vannier, B., Zhu, M.X., Brown, D., and Birnbaumer, L., The membrane topology of hTRP3 as inferred from glycosylation scanning mutagenesis and epitope immunocytochemistry, J. Biol. Chem., 273, 8675, 1998. 53. Zhu, X., Jiang, M., and Birnbaumer L., Receptor-activated Ca2+ inﬂux via human trp3 stably expressed in human embryonic kidney (HEK) 293 cells. Evidence for a non-capacitative Ca2+ entry, J. Biol. Chem., 273, 133, 1998. 54. Bloomquist, B.T., Shortridge, R.D., Schneuwly, S., Perdew, M., Montell, C., Steller, H., Rubin, G., and Pak, W.L., Isolation of a putative phospholipase C gene of Drosophila, norpA, and its role in phototransduction, Cell, 54, 723, 1988. 55. Venkatachalam, K., Zheng, F., and Gill, D.L., Regulation of canonical transient receptor potential (TRPC) channel function by diacylglycerol and protein kinase C, J. Biol. Chem., 278, 29031, 2003. 56. Shieh, B.-H. and Zhu, M.-Y., Regulation of the TRP Ca2+ channel by INAD in Drosophila photoreceptors, Neuron, 16, 991, 1996. 57. Huber, A., Sander, P., Gobert, A., Baehner, M., Hermann, R., and Paulsen, R., The transient receptor potential protein (Trp), a putative store-operated Ca2+ channel essential for phosphoinositide-mediated photoreception, forms a signaling complex with NorpA, InaC, and InaD, EMBO J., 15, 7036, 1996.

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TRP Ion Channel Function in Sensory Transduction 58. Xu, X.-Z.S., Choudhury, A., Li, X., and Montell, C., Coordination of an array of signaling proteins through homo- and heteromeric interactions between PDZ domains and target proteins, J. Cell Biol., 142, 545, 1998. 59. Chevesich, J., Kreuz, A.J., and Montell, C., Requirement for the PDZ domain protein, INAD, for localization of the TRP store-operated channel to a signaling complex, Neuron, 18, 95, 1997. 60. Montell, C., Physiology, phylogeny and functions of the TRP superfamily of cation channels, Science’s SKTE, http://skte.sciencemag.org/cgi/content/full/OC_sigtrans; 2001/90/re1, 2001. 61. Tang, Y., Tang, J., Chen, Z., Trost, C., Flockerzi, V., Li, M., Rameshi, V., and Zhu, M.X., Association of mammalian Trp4 and phospholipase C isozymes with a PDZ domain-containing protein, NHERF, J. Biol. Chem., 275, 37559, 2000. 62. Patterson, R.L., van Rossum, D.B., and Gill, D.L., Store-operated Ca2+ entry: evidence for a secretion-like coupling model, Cell, 98, 487, 1999. 63. Kim, J.Y. and Saffen, D., Activation of M1 muscarinic acetylcholine receptors stimulates the formation of a multiprotein complex centered on TRPC6 channels, J. Biol. Chem., 280, 32035, 2005. 64. Rios, E. and Pizarro, G., Voltage sensor of excitation-contraction coupling in skeletal muscle, Physiol. Rev., 71, 849, 1991. 65. Boulay, G., Brown, D.M., Qin, N., Jiang, M., Dietrich, A., Zhu, M.X., Chen, Z., Birnbaumer, M., Mikoshiba, K., and Birnbaumer, L., Modulation of Ca2+ entry by polypeptides of the inositol 1,4,5-trisphosphate receptor (IP3R) that binds transient receptor potential (TRP): evidence for roles of TRP and IP3R in store depletionactivated Ca2+ entry, Proc. Natl. Acad. Sci. USA, 96, 14955, 1999. 66. Rosado, J.A. and Sage, S.O., Activation of store-mediated calcium entry by secretionlike coupling between the inositol 1,4,5-trisphosphate receptor type II and human transient receptor potential (hTrp1) channels in human platelets, Biochem. J., 356, 191, 2001. 67. Hofmann, T., Obukhov, A.G., Scharfer, M., Harteneck, C., Gudermann, T., and Schultz, G., Direct activation of human TRPC6 and TRPC3 channels by diacyglycerol, Nature, 397, 259, 1999. 68. Trebak, M., Hempel, N., Wedel, B.J., Smyth, J.T., Bird, G.S., and Putney, J.W. Jr., Negative regulation of TRPC3 channels by protein kinase C-mediated phosphorylation of serine 712, Mol. Pharmacol., 67, 558, 2005. 69. Lucas, P., Ukhanov, K., Leinders-Zufall, T., and Zufall, F., A diacylglycerol-gated cation channel in vomeronasal neuron dendrites is impaired in TRPC2 mutant mice: mechanism of pheromone transduction, Neuron, 40, 551, 2003. 70. Cayouette, S., Lussier, M.P., Mathieu, E.L., Bousquet, S.M., and Boulay, G., Exocytotic insertion of TRPC6 channel into the plasma membrane upon Gq protein-coupled receptor activation, J. Biol. Chem., 279, 7241, 2004. 71. Bezzerides, V., Ramsey, S., Greka, A., and Clapham, D.E., Rapid translocation and insertion of TRPC5 channels, Nature Cell Biol., 6, 709, 2004. 72. Launay, P., Fleig, A., Perraud, A.L., Scharenberg, A.M., Penner, R., and Kinet, J.P., TRPM4 is a Ca2+-activated nonselective cation channel mediating cell membrane depolarization, Cell, 109, 397, 2002. 73. Soboloff, J., Spassova, M., Xu, W., He, L.P., Cuestal, N., and Gill, D.L., Role of endogenous TRPC6 channels in Ca2+ signal generation in A7r5 smooth muscle cells, J. Biol. Chem., 280, 39786, 2005.

The TRPC Family of Ion Channels

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74. Lee, K.-M., Toscas, K., and Villereal, M.L., Inhibition of bradikynin- and thapsigargin-induced calcium entry tyrosine kinase inhibitors, J. Biol. Chem., 268, 9945, 1993. 75. Lee, K.-M. and Villereal, M.L., Tyrosine phosphorylation and activation of pp60c-src and pp125FAK in bradikynin-stimulated ﬁbroblasts, Am. J. Physiol., 270, C1430, 1996. 76. Babnigg, G., Bowersox, S.R., and Villereal, M.L., Role of pp60c-src in the regulation of calcium via store-operated calcium channels, J. Biol. Chem., 272, 29434, 1997. 77. Gutkind, J.S. and Robbins, K.C., Activation of transforming G-protein–coupled receptors induces rapid tyrosine phosphorylation of cellular proteins including p125FAK and p130src substrate, Biochem. Biophys. Res. Commun., 188, 155, 1992. 78. Igishi, T. and Gutkind, J.S., Tyrosine kinases of the src family participate in signaling to MAP kinase from both Gq- and Gi-coupled receptors, Biochem. Biophys. Res. Commun., 244, 5, 1998. 79. Vazquez, G., Wedel, B.J., Kawasaki, B.T., St. John Bird, G., and Putney, J., Obligatory role for src kinase in the signaling mechanism for TRPC3 cation channels, J. Biol. Chem., 279, 40521, 2004. 80. Kawasaki, B.T., Liao, Y., and Birnbaumer, L., Role of Src in C3 transient receptor potential channel function and evidence for a heterogeneous makeup of receptor- and store-operated Ca2+ entry channels, Proc. Natl. Acad. Sci. USA, 103, 335, 2006. 81. Hisatsune, C., Kuroda, Y., Nakamura, K., Inoue, T., Nakamura, T., Michikawa, T., Mitsutani, A., and Mikoshiba K., Regulation of TRPC6 channel activity by tyrosine phosphorylation, J. Biol. Chem., 279, 18887, 2004. 82. Zagranichnaya, T.K., Wu, X., and Villereal, M.L., Endogenous TRPC1, TRPC3, and TRPC7 proteins combine to form native store-operated channels in HEK-293 cells, J. Biol. Chem., 280, 29559, 2005. 83. Freichel, M., Vennekens, R., Olausson, J., Hoffmann, M., Muller, C., Stolz, S., Scheunemann, J., Weissgerber, P., and Flockerzi, V., Functional role of TRPC proteins in vivo: lessons from TRPC4-deﬁcient mouse models, Biochem. Biophys. Res. Commun., 322, 1352, 2004. 84. Dietrich, A., Mederos y Schnitzler, M., Emmel, J., Kalwa, H., Hofmann, T., and Gudermann, T., N-linked protein glycosylation is a major determinant for basal TRPC3 and TRPC6 channel activity, J. Biol. Chem., 278, 47842, 2003. 85. Rosker, C., Graziani, A., Lukas, M., Eder, P., Zhu, M.X., Romanin, C., and Groschner, K., Ca2+ signaling by TRPC3 involves Na+ entry and local coupling to the Na+/Ca2+ exchanger, J. Biol. Chem., 279, 13696, 2004. 86. Schindl, R., Kahr, H., Graz, I., Groschner, K., and Romanin, C., Store depletionactivated CaT1 currents in rat basophilic leukemia mast cells are inhibited by 2aminoethoxydiphenyl borate. Evidence for a regulatory component that controls activation of both CaT1 and CRAC (Ca2+ release-activated Ca2+ channel) channels, J. Biol. Chem., 277, 26950, 2002. 87. Zhang, Z., Tang, T., Tikunova, S., Johnson, J.D., Chen, Z., Qin, N., Dietrich, A., Stefani, E., Birnbaumer, L., and Zhu, M.X., Activation of TRP3 by IP3 receptor through displacement of inhibitory calmodulin from a common binding site, Proc. Natl. Acad. Sci. USA, 98, 3168, 2001. 88. Tang, J., Lin, Y., Zhang, Z., Tikunova, S., Birnbaumer, L., and Zhu, M.X., Identiﬁcation of common binding sites for calmodulin and IP3 receptors on the carboxyltermini of TRP channels, J. Biol. Chem., 276, 21303, 2001.

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89. Trost, C., Bergs, C., Himmerkus, N., and Flockerzi, V., The transient receptor potential, TRP4, cation channel is a novel member of the family of calmodulin-binding proteins, Biochem. J., 355, 663, 2001. 90. Yildirim, E., Dietrich, A., and Birnbaumer, L., Characterization of mouse TRPC2 transcripts: alternative splicing of the primary transcript and calmodulin binding to the translated N-terminus, Proc. Natl. Acad. Sci. USA, 100, 2220, 2003. 91. Liu, X., Wang, W., Singh, B.B., Lockwich, T., Jadlowiec, J., O’Connell, B., Wellner, R., Zhu, M.X., and Ambudkar, I.S., Trp1, a candidate protein for the store-operated Ca2+ inﬂux mechanism in salivary gland cells, J. Biol. Chem., 275, 3403, 2000. 92. Vannier, B., Peyton, M., Boulay, G., Brown, D., Qin, N., Jiang, M., Zhu, X., and Birnbaumer, L., Mouse trp2, the homologue of the human trpc2 pseudogene, encodes mTrp2, a store depletion-activated capactitative Ca2+ entry channel, Proc. Natl. Acad. Sci. USA, 96, 2060, 1999. 93. Jungnickel, M.S., Marrero, H., Birnbaumer, L., Lemos, J.R., and Florman, H.M., Trp2 regulates Ca2+ entry into sperm triggered by egg ZP3, Nature Cell Biol., 3, 499, 2001. 94. Vazquez, G., Wedel, B.J., Trebak, M., Bird, G.S., and Putney, J.W. Jr., Expression level of the canonical transient receptor potential 3 (TRPC3) channel determines its mechanism of activation, J. Biol. Chem., 278, 21649, 2003. 95. Yildirim, E., Kawasaki, B.T., and Birnbaumer, L., Molecular cloning of TRPC3a, an N-terminally extended, store-operated variant of the human C3 transient receptor potential channel, Proc. Natl. Acad. Sci. USA, 102, 3307, 2005. 96. Philipp, S., Cavalie, A., Freichel, M., Wissenbach, U., Zimmer, S., Trost, C., Marquart, A., Murakami, M., and Flockerzi, V., A mammalian capacitative calcium entry channel homologous to Drosophila TRP and TRPL, EMBO J., 15, 6166, 1996. 97. Philipp, S., Hambrecht, J., Braslavski, L., Schroth, G., Freichel, M., Murakami, M., Cavalie, A., and Flockerzi, V., A novel capacitative calcium entry channel expressed in excitable cells, EMBO J., 17, 4274, 1998. 98. Riccio, A., Mattei, C., Kelsell, R.E., Medhurst, A.D., Calver, A.R., Randall, A.D., Davis, J.D., Benham, C.D., and Pangalos, M.N., Cloning and functional expression of human short TRP7, a candidate protein for store-operated Ca2+ inﬂux, J. Biol. Chem., 277, 12302, 2002. 99. Lievremont, J.P., Bird, G.S., and Putney, J.W. Jr., Canonical transient receptor potential TRPC7 can function as both a receptor and store-operated channel in HEK-293 cells, Am. J. Physiol. Cell Physiol., 287, C1709, 2004. 100. Schaefer, M., Plant, T.D., Stresow, N., Albrecht, N., and Schultz, G., Functional differences between TRPC4 splice variants, J. Biol. Chem., 277, 3752–3759, 2002. 101. Jung, S., Muehle, A., Schaefer, M., Strotmann, R., Schultz, G., and Plant, T.D., Lanthanides potentiate TRPC5 currents by an action at extracellular sites close to the pore mouth, J. Biol. Chem., 278, 3562, 2003. 102. Inoue, R., Okada, T., Onoue, H., Hara, Y., Shimizu, S., Naitoh, S., Ito, Y., and Mori, Y., The transient receptor potential protein homologue TRP6 is the essential component of vascular 1-adrenoceptor-activated Ca2+-permeable cation channel, Circ. Res., 88, 325, 2001.

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Multiple Mechanisms of TRPC Activation James W. Putney, Jr. National Institute of Environmental Health Sciences

CONTENTS Abstract ....................................................................................................................31 Introduction..............................................................................................................31 TRPCs: Mechanisms of Regulation in Transfecto..................................................32 Mechanisms of TRPC Activation in Situ ................................................................35 Summary ..................................................................................................................36 Acknowledgments....................................................................................................37 References................................................................................................................37

ABSTRACT TRPC (canonical transient receptor potential) channels are vertebrate homologues of the Drosophila photoreceptor channel (TRP). Considerable research has been brought to bear on the seven members of this family, especially with regard to their possible roles in calcium entry. Unfortunately, the current literature presents a confusing picture, with different laboratories describing widely differing results and interpretations. It appears that ectopically expressed TRPC channels can be activated as a consequence of phospholipase C activation (by increases in diacylglycerols or by loss of phosphatidylinositol 4,5-bisphosphate), by stimulation of trafﬁcking to the plasma membrane, or by depletion of intracellular Ca2+ stores. These diverse experimental ﬁndings arise because TRPC channels can, under both experimental as well as physiological conditions, be activated in three distinct ways, possibly depending on their subunit composition or signaling complex environment. The TRPCs may be unique among ion channel subunit families in having the ability to participate in the assembly and function of multiple types of physiologically important ion channels.

INTRODUCTION The mammalian TRPC genes were identiﬁed from searches for homologues of the Drosophila trp gene.1–10 The proteins encoded by the TRPC genes, like the parental Drosophila TRP protein, span the membrane six times11 and contain a short 31

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hydrophobic sequence believed to be involved in forming the pore of the channel.12 By analogy with other similar channels, it is believed that a functional TRPC channel will be formed by the four TRPC proteins coming together12; thus channels could be formed as homotetramers, if all four TRPCs are the same, or heterotetramers, if more than one kind of TRPC is involved. There appear to be restrictions as to which TRPCs can come together to form channels, although there is not complete agreement on what these restrictions are.12,13 Hofmann et al.12 found that when TRPC channels were exogenously expressed in HEK293 cells, the only permitted combinations were TRPC3, 6, and 7, or TRPC1, 4, and 5. However, a number of reports have found that TRPC1 and 3 can associate, with both exogenous and endogenous expressions.13–15 Unlike previous studies of ion channel function, investigators attempted to understand the function of mammalian TRPC channels beginning only with knowledge of their coding sequence. There was considerable interest in this group because of the suspicion that they might be the long-sought store-operated or capacitative calcium entry channels.16 In addition to TRPCs, the TRP superfamily includes two other major families: TRPMs (melastatin-related transient receptor potential channels), related structurally to melastatin, and TRPVs (vanilloid receptor-related transient potential channels), related to the vanilloid receptor, as well as more distantly related genes identiﬁed in genetic screens for speciﬁc diseases and whose physiological functions are less certain. Considerable progress has been made in our understanding of a number of TRP channels, especially those in the TRPV and TRPM families,17,18 and much of this information is reviewed in other chapters in this book. However, the function and regulation of TRPC channels have been more problematic, plagued with conﬂicting ﬁndings and interpretations. In this review I attempt to organize and rationalize some of these problems. I propose a paradigm that can help to explain the somewhat poorly predictable behavior of these widely expressed cation channels. In addition, I suggest that the apparently poorly predictable behavior of these channels in laboratory experiments might reﬂect expression of at least three distinct modes of regulation in their native physiological environments. A number of other aspects of TRPC channel regulation and activation mechanisms are reviewed in chapters by Birnbaumer, Gudermann, and Groschner.

TRPCS: MECHANISMS OF REGULATION IN TRANSFECTO Based on sequence similarities, the seven TRPCs can be conveniently divided into four subgroups: TRPC1, TRPC2, and two additional groups, TRPC3, 6, and 7, and TRPC4 and 5. TRPC1 is most closely related to TRPC4 and 5 and is sometimes considered to be in the same group. TRPC2 is a pseudogene in humans but not in other mammalian species; there is evidence that TRPC2 can function as a storeoperated channel in sperm,8,19 and is likely involved in vomeronasal function.20,21 The body of literature on this channel is small by comparison to other TRPCs, however, and I will not discuss it further here. This topic is covered in the chapter by Dulac and Liman. The majority of research dealing with TRPCs has focused on

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the TRPC1/4/5 or TRPC3/6/7 groups. To date there is no clear evidence that TRPC5 behaves in a fundamentally different manner from TRPC4, and these two channels appear capable of association with TRPC1. Likewise, when compared side by side, TRPC3, 6, and 7 seem to behave similarly. There are important exceptions to these generalities, however, as noted below. The TRPCs are so named22 because this family is most closely related to the original Drosophila channel TRP (i.e., they are the “canonical” TRPs). Because Drosophila TRP was clearly activated downstream of phospholipase C and inositol 1,4,5-trisphosphate (IP3) generation in photoreceptor cells, Hardie and Minke23 suggested that TRP might be the long-sought store-operated Ca2+ channel. The store-operated or capacitative calcium entry channels are activated by a process triggered by the depletion of intracellular endoplasmic reticulum Ca2+ stores.16,24 Recent ﬁndings indicate that the sensor of intracellular Ca2+ store depletion may be the Ca2+-binding membrane protein, STIM1.25,26 Ironically, it is now very clear that store depletion is not the mechanism by which Drosophila TRP is activated in its native environment in photoreceptor cells.27 The signiﬁcance of this basic ﬁnding to the larger body of work on mammalian TRPCs perhaps should not be overlooked. Yet it is clear from phospholipase C (PLC)-deﬁcient mutants that Drosophila TRP is activated in some manner downstream of PLC. It is most likely, although not proven, that a lipid mediator derived from PLC products activates TRP, perhaps the immediate breakdown product diacylglycerol (DAG), and it is also regulated in complex ways by the substrate of PLC, phosphatidylinositol 4,5bisphosphate (PIP2).27 For the seven mammalian TRPCs, there is evidence that these channels are also activated in some manner downstream of PLC. Although initial studies suggested that TRPC1 and 3 were activated by store depletion,3 it was subsequently shown that activation resulted from the constitutive activity of the channels28 (however, see reference 29). One report suggested that IP3 and the IP3 receptor were involved in activating TRPC3,30 but other laboratories failed to reproduce these results.7,31 Likewise, the original studies on TRPC4 and TRPC5 presented evidence for activation by store depletion,5,6 but other researchers failed to reproduce these ﬁndings32,33 (but see reference 34). For the TRPC3/6/7 subfamily, the vast majority of published results suggests that DAG, produced upon phospholipase C activation, is the signal activating these channels.7,9,15,31,35–38 For TRPC4 and 5, the case is not so clear. TRPC4 and 5 are highly sensitive to inhibition by protein kinase C, making it difﬁcult to obtain evidence that DAG can activate. Indeed, while TRPC3/6/7 channels can be activated by exogenous DAG (usually oleyl acetyl glycerol, or OAG), OAG inhibits the activation of TRPC4 or 5, and this inhibition is blocked by inhibitors of protein kinase C.36 Thus, if DAG is involved in the activation of TRPC4 or 5, there must be considerable compartmentalization of the signaling pathway to prevent concomitant inhibition by protein kinase C. An interesting possibility is that TRPC4 and 5 may be tonically inhibited by PIP2, such that PLC-mediated degradation of PIP2 relieves this inhibition, resulting in channel activation. Normally, PIP2 is thought to provide a positive regulation of TRPs39–41; however, for TRPV1, PIP2 appears to regulate negatively, although this is not thought to constitute a primary mechanism for channel activation.42

34

TRP Ion Channel Function in Sensory Transduction

As mentioned above, TRPC1 is often considered with TRPC4 and 5, as it is somewhat similar in sequence and is known to associate with TRPC4 and 5 to form heteromultimeric channels.12,13 The actions of TRPC1 when expressed on its own are controversial; some researchers demonstrate activated ion channel behavior following expression of TRPC1,3,4,43–45 while others ﬁnd that the channel does not trafﬁc to the plasma membrane correctly unless coexpressed with TRPC4 or 5.12 Interestingly, and important for arguments to be advanced below, Strübing et al.46 found that heteromultimers of TRPC1 and TRPC5 had different electrophysiological properties from homotetramers of TRPC5. A second mechanism that is important for activating TRPC channels involves regulation of their trafﬁcking to the plasma membrane.47–49 The published reports on this topic are not consistent with one another. However, it is clear from the study of Bezzerides et al.49 that growth factors (EGF) activate the insertion of TRPC5 channels into the plasma membrane. Once the channels reach the membrane, subsequent maneuvers that activate the channels result in larger currents. Reports from other laboratories indicate that for other TRPCs, the translocation event per se leads to increased currents.47,48 In either case, either constitutive or stimulated currents would be increased simply by increasing the number of channels and would not necessarily involve any further increase in open probability. The mechanism by which TRPC5 secretion is signaled does not seem to involve activation of PLC; instead it involves a pathway requiring phosphoinositide 3-kinase, Rac1, and phosphatidylinositol 4-kinase.49 Finally, it is signiﬁcant that only TRPC5 homotetrameric channels were activated in this manner. Heterotetramers containing TRPC5 and 1 did not translocate to the plasma membrane in response to EGF. This mechanism of regulation may apply to members of the broader TRP superfamily; there is at least one example of a TRPV channel that appears to be regulated in this way.50 The third major mechanism that has been described for TRPC channel activation is the store-depletion or capacitative calcium entry mechanism. A number of studies have reported activation of TRPC channels by store depletion,3–6,30,38,51–61 and knockout or knock-down of TRPCs often reduces store-operated calcium entry.44,61–69 Yet in many instances these channels clearly are not store operated.7,9,31,32,35,37,60,70,71 It is becoming increasingly clear that the basis for the different behaviors is expression conditions. In DT40 B-lymphocytes, TRPC3 formed a store-operated channel at low levels of expression and formed a DAG-activated channel at higher levels of expression.60 The loss of store-operated behavior at higher expression levels may result from inappropriate stoichiometry among members of a signaling complex,10 as has been argued previously for scaffolding proteins.72,73 Signiﬁcantly, the pharmacology of the channels differed in these two modes; the store-operated TRPC3 channels were much more sensitive to block by Gd3+ than the nonstore-operated channels.59 Both TRPC738 and TRPC534 are store operated when stably expressed in HEK293 cells; in this case, the store-operated channels are either capable of activation by alternative, nonstore-operated mechanisms, or they coexist with channels that are activated by the nonstore-operated mechanisms. For TRPC7, it was demonstrated that this store-operated behavior is only seen with stable transfection. Transient transfection of HEK293 cells with TRPC7 results in channels that can only be activated by receptor activation or OAG, not by store depletion.38

Multiple Mechanisms of TRPC Activation

35

Thus, the available data demonstrate that, when ectopically expressed, TRPCs can be activated by one of three mechanisms: store depletion, phospholipase C (or its products), or channel translocation to the plasma membrane. The mode of activation seems to depend on the environment in which the subunit ﬁnds itself, including perhaps the nature of other subunits that compose the channel pore. This principle is most clearly illustrated by the case of translocation of TRPC5, which only occurs with TRPC5 homotetramers, not with TRPC5/1 heterotetramers.49 It is also a logical interpretation of the observed difference in Gd3+ sensitivity of TRPC3 channels in store-operated as compared to phospholipase C-activated mode.59 The question then arises: do these different modes of regulation occur with TRPC subunits when expressed in their native environments in untransfected cells?

MECHANISMS OF TRPC ACTIVATION IN SITU The answer to that question seems to be, yes. There is now evidence that each of these modes of TRPC function in native, untransfected cells. In the case of storeoperated channels, I have already mentioned the considerable number of examples of disrupted behavior of native store-operated channels by knock-down of TRPCs. However, a sticking point has been the failure of ectopically expressed TRPCs (or as yet any expressed ion channel) to faithfully reproduce the electrophysiological characteristics of Icrac (ﬁrst described in detail in references 74 and 75; for an example, see also reference 76). However, it is clear that there are a variety of different kinds of store-operated channels,77–84 some of which have properties reminiscent of TRPCs. It is realistic to expect that TRPCs may contribute to the composition of store-operated channels in such instances. It is not out of the question to consider that TRPCs might play a role in Icrac channels, given the possibility of heteromultimers with drastically altered electrophysiological properties. Two reports have provided evidence for a role for TRPC168 and TRPC385 in Icrac. There are examples of native channels that appear to be activated by diacylglycerols under physiological conditions.70,71,86–90 In a particularly thorough study, Inoue et al.87 investigated the role of TRPC6 in α-adrenergic-activated cation channels. These authors examined the electrophysiological and pharmacological characteristics of the endogenous cation entry controlled by α1-adrenoceptors in rabbit portal vein smooth muscle cells and found this cation entry to be reminiscent of that of the TRPC6 current when transiently expressed in HEK293 cells. Both the endogenous current and the current due to expressed TRPC6 could be activated by OAG in a PKC-independent fashion. Furthermore, TRPC6 mRNA and protein expression in the smooth muscle cells and knock-down of TRPC6 with antisense RNA in smooth muscle cells resulted in almost complete abrogation of the α1-adrenergic response. Similarly, Jung et al.71 showed that vasopressin activates a non-CCE, nonselective cation current in the smooth muscle cell line, A7r5, resembling that seen with expression of TRPC6 in HEK293 cells. The DAG analog, OAG, activated the endogenous, agonist-sensitive current in A7r5 cells, and those cells were shown to express mRNA for TRPC1 and TRPC6 but not for other TRPC proteins. The authors then concluded that TRPC6 is likely a component of the endogenous agonistactivated channel in the A7r5 cell line.

36

TRP Ion Channel Function in Sensory Transduction

There are far fewer examples of stimulated translocation of native TRPC channels, likely due to the rather recent discovery of this mode of activation. However, Bezzerides et al.49 reported that native TRPC5 channels were localized to active growth cones in cultured hippocampal neurons (see also reference 91). Surface expression of TRPC5 was increased by several different growth factors. TRPC1 was expressed in the cell soma and processes, and not in growth cones, consistent with the observation that only TRPC5 homotetramers undergo regulation through translocation.

SUMMARY I previously suggested that TRPCs could form distinct types of channels, being regulated by either phospholipase C-dependent or store-depletion mechanisms.10 Now the number of possible modes of regulation has expanded to three92 (see Figure 2.1), and we cannot know that others will not be discovered. It is also becoming

FIGURE 2.1 Activation mechanisms for calcium-permeable TRPC cation channels. TRPC channels can be activated in any of three distinct ways. (From left) Channels sequestered in a vesicular compartment can be translocated to the plasma membrane in response to growth factor (GF) whence the expression of their constitutive activity will contribute to membrane signaling and electrical properties. TRPC channels can be activated by DAG, or possibly as a result of loss of PIP2 (not shown) as a result of agonist (Ag) activation of phospholipase C (PLC) by a G-protein-coupled (G) pathway. In some instances, this activation mode requires the tyrosine kinase, Src,93 and is negatively regulated by protein kinase C (PKC).9,31,36,94,95 Finally, the activation of PLC leads to the production of IP3, which activates the IP3 receptor (IP3R) and causes the release of Ca2+ from a critical component of the endoplasmic reticulum. This can in turn activate TRPC channels through the capacitative or store-operated pathway, possibly involving the Ca2+ sensor protein, STIM1. There is evidence that in the latter case, the subunit composition of the store-operated channels may differ from that of the PLCregulated channels.

Multiple Mechanisms of TRPC Activation

37

increasingly clear that this multiplicity of regulatory mechanisms does not simply reﬂect (at least in all cases) aberrant behavior due to overexpression but rather is indicative of true diversity of channel function in vivo. At least one of the factors that determines the function and regulation of TRPC channels appears to be the subunit composition of the assembled tetrameric channel. Additional factors may include partners in a signaling complex, such as regulatory subunits or scaffolding structures. The propensity of TRPC channels to intereact with the Ca2+ sensor, STIM1,25,26 may be important for their regulation by Ca2+ store depletion. As argued previously,10 the ability of cells to utilize TRPCs in diverse ways may have significance beyond the ion channel ﬁeld; such a multiplicity offers a means by which the complex mammalian organism can be assembled from what has turned out to be a surprisingly limited genome.

ACKNOWLEDGMENTS The author gratefully acknowledges ideas and criticisms from Mohamed Trebak, Christian Erxleben, and Steve Shears.

REFERENCES 1. Zhu, X., Chu, P.B., Peyton, M., and Birnbaumer L., Molecular cloning of a widely expressed human homologue for the Drosophila trp gene, FEBS Letters 373, 193, 1995. 2. Wes, P.D., Chevesich, J., Jeromin, A., Rosenberg, C., Stetten, G., and Montell, C., TRPC1, a human homolog of a Drosophila store-operated channel, Proc. Nat. Acad. Sci. USA 92, 9652, 1995. 3. Zhu, X., Jiang, M., Peyton, M., Boulay, G., Hurst, R., Stefani, E., and Birnbaumer, L., trp, a novel mammalian gene family essential for agonist-activated capacitative Ca2+ entry, Cell 85, 661, 1996. 4. Zitt, C., Zobel, A., Obukhov, A.G., Harteneck, C., Kalkbrenner, F., Lückhoff, A., and Schultz, G., Cloning and functional expression of a human Ca2+-permeable cation channel activated by calcium store depletion, Neuron 16, 1189, 1996. 5. Philipp, S., Cavalié, A., Freichel, M., Wissenbach, U., Zimmer, S., Trost, C., Marguart, A., Murakami, M., and Flockerzi, V., A mammalian capacitative calcium entry channel homologous to Drosophila TRP and TRPL, EMBO J. 15, 6166, 1996. 6. Philipp, S., Hambrecht, J., Braslavski, L., Schroth, G., Freichel, M., Murakami, M., Cavalié, A., and Flockerzi, V., A novel capacitative calcium entry channel expressed in excitable cells, EMBO J. 17, 4274, 1998. 7. Hofmann, T., Obukhov, A.G., Schaefer, M., Harteneck, C., Gudermann, T., and Schultz, G., Direct activation of human TRPC6 and TRPC3 channels by diacylglycerol, Nature 397, 259, 1999. 8. Vannier, B., Peyton, M., Boulay, G., Brown, D., Qin, N., Jiang, M., Zhu, X., and Birnbaumer, L., Mouse trp2, the homologue of the human trpc2 pseudogene, encodes mTrp2, a store depletion–activated capacitative Ca2+ channel, Proc. Nat. Acad. Sci. USA 96, 2060, 1999. 9. Okada, T., Inoue, R., Yamazaki, K., Maeda, A., Kurosaki, T., Yamakuni, T., Tanaka, I., Shimizu, S., Ikenaka, K., Imoto, K., and Mori, Y., Molecular and functional characterization of a novel mouse transient receptor potential protein homologue

38

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10. 11.

12.

13.

14. 15.

16. 17.

18. 19.

20. 21.

22.

23.

24. 25.

26.

TRP7. Ca2+-permeable cation channel that is constitutively activated and enhanced by stimulation of G protein-coupled receptor, J. Biol. Chem. 274, 27359, 1999. Putney, J.W., Jr., The enigmatic TRPCs: multifunctional cation channels, Trends Cell Biol. 14, 282, 2004. Vannier, B., Zhu, X., Brown, D., and Birnbaumer, L., The membrane topology of human transient receptor potential 3 as inferred from glycosylation-scanning mutagenesis and epitope immunocytochemistry, J. Biol. Chem. 273, 8675, 1998. Hofmann, T., Schaefer, M., Schultz, G., and Gudermann, T., Subunit composition of mammalian transient receptor potential channels in living cells, Proc. Nat. Acad. Sci. USA 99, 7461, 2002. Strübing, C., Krapivinsky, G., Krapivinsky, L., and Clapham, D.E., Formation of novel TRPC channels by complex subunit interactions in embryonic brain, J. Biol. Chem. 278, 39014, 2003. Xu, X.-Z.S., Li, H.-S., Guggino, W.B., and Montell, C., Coassembly of TRP and TRPL produces a distinct store-operated conductance, Cell 89, 1155, 1997. Lintschinger, B., Balzer-Geldsetzer, M., Baskaran, T., Graier, W.F., Romanin, C., Zhu, M.X., and Groschner, K., Coassembly of Trp1 and Trp3 proteins generates diacylglycerol- and Ca2+-sensitive cation channels, J. Biol. Chem. 275, 27799, 2000. Putney, J.W., Jr., Capacitative Calcium Entry, Landes Biomedical Publishing, Austin, TX, 1997. Gunthorpe, M.J., Benham, C.D., Randall, A., and Davis, J.B., The diversity in the vanilloid (TRPV) receptor family of ion channels, Trends Pharmacol. Sci. 23, 183, 2002. Clapham, D.E., TRP channels as cellular sensors, Nature 426, 517, 2003. Jungnickel, M.K., Marreo, H., Birnbaumer, L., Lémos, J.R., and Florman, H.M., Trp2 regulates entry of Ca2+ into mouse sperm triggered by egg ZP3, Nature Cell Biol. 3, 499, 2001. Liman, E.R., Corey, D.P., and Dulac, C., TRP2: A candidate transduction channel for mammalian pheromone sensory signaling, Proc. Nat. Acad. Sci. USA 96, 5791, 1999. Stowers, L., Holy, T.E., Meister, M., Dulac, C., and Koentges, G., Loss of sex discrimination and male-male aggression in mice deﬁcient for TRP2, Science 295, 1493, 2002. Montell, C., Birnbaumer, L., Flockerzi, V., Bindels, R.J., Bruford, E.A., Caterina, M.J., Clapham, D.E., Harteneck, C., Heller, S., Julius, D., Kojima, I., Mori, Y., Penner, R., Prawitt, D., Scharenberg, A.M., Schultz, G., Shimizu, N., and Zhu, M.X., A uniﬁed nomenclature for the superfamily of TRP cation channels, Mol. Cell 9, 229, 2002. Hardie, R.C. and Minke, B., Novel Ca2+ channels underlying transduction in Drosophila photoreceptors: implications for phosphoinositide-mediated Ca2+ mobilization, Trends Neurosci. 16, 371, 1993. Parekh, A.B. and Penner, R., Store depletion and calcium inﬂux, Physiol. Rev. 77, 901, 1997. Roos, J., DiGregorio, P.J., Yeromin, A.V., Ohlsen, K., Lioudyno, M., Zhang, S., Safrina, O., Kozak, J.A., Wagner, S.L., Cahalan, M.D., Velicelebi, G., and Stauderman, K.A., STIM1, an essential and conserved component of store-operated Ca2+ channel function, J. Cell Biol. 169, 435, 2005. Liou, J., Kim, M.L., Heo, W.D., Jones, J.T., Myers, J.W., Ferrell, J.E., Jr., and Meyer, T., STIM is a Ca2+ sensor essential for Ca2+-store-depletion–triggered Ca2+ inﬂux, Curr. Biol. 15, 1235, 2005.

Multiple Mechanisms of TRPC Activation

39

27. Hardie R.C., Regulation of trp channels via lipid second messengers, Ann. Rev. Physiol. 65, 735, 2003. 28. Zhu, X., Jiang, M., and Birnbaumer, L., Receptor-activated Ca2+ inﬂux via human Trp3 stably expressed in human embryonic kidney (HEK)293 cells. Evidence for a non-capacitative calcium entry, J. Biol. Chem. 273, 133, 1998. 29. Yildirim, E., Kawasaki, B.T., and Birnbaumer, L., Molecular cloning of TRPC3a, an N-terminally extended, store-operated variant of the human C3 transient receptor potential channel, Proc. Natl. Acad. Sci USA 102, 3307, 2005. 30. Kiselyov, K., Xu, X., Mozhayeva, G., Kuo, T., Pessah, I., Mignery, G., Zhu, X., Birnbaumer, L., and Muallem, S., Functional interaction between InsP3 receptors and store-operated Htrp3 channels, Nature 396, 478, 1998. 31. Trebak, M., Bird, G.St.J., McKay, R.R., Birnbaumer, L., and Putney, J.W., Jr., Signaling mechanism for receptor-activated TRPC3 channels, J. Biol. Chem. 278, 16244, 2003. 32. Schaefer, M., Plant, T.D., Obukhov, A.G., Hofmann, T., Gudermann, T., and Schultz, G., Receptor-mediated regulation of the nonselective cation channels TRPC4 and TRPC5, J. Biol. Chem. 275, 17517, 2000. 33. McKay, R.R., Szmeczek-Seay, C.L., Lièvremont, J.-P., Bird, G.St.J., Zitt, C., Jüngling, E., Lückhoff, A., and Putney, J.W., Jr., Cloning and expression of the human transient receptor potential 4 (TRP4) gene: localization and functional expression of human TRP4 and TRP3, Biochem. J. 351, 735, 2000. 34. Zeng, F., Xu, S.Z., Jackson, P.K., McHugh, D., Kumar, B., Fountain, S.J., and Beech, D.J., Human TRPC5 channel activated by a multiplicity of signals in a single cell, J. Physiol. 559, 739, 2004. 35. Trebak, M., Vazquez, G., Bird, G.St.J., and Putney, J.W., Jr., The TRPC3/6/7 subfamily of cation channels, Cell Calcium 33, 451, 2003. 36. Venkatachalam, K., Zheng, F., and Gill, D.L., Regulation of canonical transient receptor potential (TRPC) channel function by diacylglycerol and protein kinase C, J Biol. Chem. 278, 29031, 2003. 37. Zhang, L. and Saffen, D., Muscarinic acetylcholine receptor regulation of TRP6 Ca2+ channel isoforms, J. Biol. Chem. 276, 13331, 2001. 38. Lièvremont, J.P., Bird, G.S., and Putney, J.W., Jr., Canonical transient receptor potential TRPC7 can function as both a receptor- and store-operated channel in HEK-293 cells, Am. J. Physiol. Cell Physiol. 287, C1709, 2004. 39. Runnels, L.W., Yue, L., and Clapham, D.E., The TRPM7 channel is inactivated by PIP2 hydrolysis, Nature Cell Biol. 4, 329, 2002. 40. Liu, B. and Qin, F., Functional control of cold- and menthol-sensitive TRPM8 ion channels by phosphatidylinositol 4,5-bisphosphate, J. Neurosci. 25, 1674, 2005. 41. Rohacs, T., Lopes, C.M., Michailidis, I., and Logothetis, D.E., PI(4,5)P2 regulates the activation and desensitization of TRPM8 channels through the TRP domain, Nat. Neurosci. 8, 626, 2005. 42. Liu, B., Zhang, C., and Qin, F., Functional recovery from desensitization of vanilloid receptor TRPV1 requires resynthesis of phosphatidylinositol 4,5-bisphosphate, J. Neurosci. 25, 4835, 2005. 43. Sinkins, W.G., Estacion, M., and Schilling, W.P., Functional expression of TrpC1: a human homologue of the Drosophila Trp channel, Biochem. J. 331, 331, 1998. 44. Liu, X., Wang, W., Singh, B.B., Lockwich, T., Jadlowiec, J., O’Connell, B., Wellner, R., Zhu, M.X., and Ambudkar, I.S., Trp1, a candidate protein for the store-operated Ca2+ inﬂux mechanism in salivary gland cells, J. Biol. Chem. 275, 3403, 2000.

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TRP Ion Channel Function in Sensory Transduction 45. Chen, J. and Barritt, G.J., Evidence that TRPC1 (transient receptor potential canonical 1) forms a Ca(2+)-permeable channel linked to the regulation of cell volume in liver cells obtained using small interfering RNA targeted against TRPC1, Biochem. J. 373, 327, 2003. 46. Strübing, C., Krapivinsky, G., Krapivinsky, L., and Clapham, D.E., TRPC1 and TRPC5 form a novel cation channel in mammalian brain, Neuron 29, 645, 2001. 47. Cayouette, S., Lussier, M.P., Mathieu, E.L., Bousquet, S.M., and Boulay, G., Exocytotic insertion of TRPC6 channel into the plasma membrane upon Gq protein-coupled receptor activation, J. Biol. Chem. 279, 7241, 2004. 48. Singh, B.B., Lockwich, T.P., Bandyopadhyay, B.C., Liu, X., Bollimuntha, S., Brazer, S.C., Combs, C., Das, S., Leenders, A.G., Sheng, Z.H., Knepper, M.A., Ambudkar, S.V., and Ambudkar, I.S., VAMP2-dependent exocytosis regulates plasma membrane insertion of TRPC3 channels and contributes to agonist-stimulated Ca(2+) inﬂux, Mol. Cell 15, 635, 2004. 49. Bezzerides, V.J., Ramsey, I.S., Kotecha, S., Greka, A., and Clapham, D.E., Rapid vesicular translocation and insertion of TRP channels, Nat. Cell Biol. 6, 709, 2004. 50. Kanzaki, M., Zhang, Y.-Q., Mashima, H., Li, L., Shibata, H., and Kojima, I., Translocation of a calcium-permeable cation channel induced by insulin-like growth factor-I, Nature Cell Biol. 1, 165, 1999. 51. Vaca, L., Sinkins, W.G., Hu, Y., Kunze, D.L., and Schilling, W.P., Activation of recombinant trp by thapsigargin in Sf9 insect cells, Am. J. Physiol. 267, C1501, 1994. 52. Birnbaumer, L., Zhu, X., Jiang, M., Boulay, G., Peyton, M., Vannier, B., Brown, D., Platano, D., Sadeghi, H., Stefani, E., and Birnbaumer, M., On the molecular basis and regulation of cellular capacitative calcium entry: roles for Trp proteins, Proc. Nat. Acad. Sci. USA 93, 15195, 1996. 53. Tomita, Y., Kaneko, S., Funayama, M., Kondo, H., Satoh, M., and Akaike, A., Intracellular Ca2+ store-operated inﬂux of Ca2+ through TRP-R, a rat homolog of TRP, expressed in Xenopus oocytes, Neurosci. Letters 248, 195, 1998. 54. Groschner, K., Hingel, S., Lintschinger, B., Balzer, M., Romanin, C., Zhu, X., and Schreibmayer, W., Trp proteins form store-operated cation channels in human vascular endothelial cells, FEBS Lett. 437, 101, 1998. 55. Kiselyov, K., Mignery, G.A., Zhu, M.X., and Muallem, S., The N-terminal domain of the IP3 receptor gates store-operated hTrp3 channels, Mol. Cell 4, 423, 1999. 56. Kinoshita, M., Akaike, A., Satoh, M., and Kaneko, S., Positive regulation of capacitative Ca2+ entry by intracellular Ca2+ in Xenopus oocytes expressing rat TRP4, Cell Calcium 28, 151, 2000. 57. Vazquez, G., Lièvremont, J.-P., Bird, G.St.J., and Putney, J.W., Jr., Human Trp3 forms both inositol trisphosphate receptor-dependent and receptor-independent storeoperated cation channels in DT40 avian B-lymphocytes, Proc. Nat. Acad. Sci. USA 98, 11777, 2001. 58. Riccio, A., Mattei, C., Kelsell, R.E., Medhurst, A.D., Calver, A.R., Randall, A.D., Davis, J.B., Benham, C.D., and Pangalos, M.N., Cloning and functional expression of human short TRP7, a candidate protein for store-operated Ca2+ inﬂux, J. Biol. Chem. 277, 12302, 2002. 59. Trebak, M., Bird, G.St.J., McKay, R.R., and Putney, J.W., Jr., Comparison of human TRPC3 channels in receptor-activated and store-operated modes. Differential sensitivity to channel blockers suggests fundamental differences in channel composition, J. Biol. Chem. 277, 21617, 2002.

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60. Vazquez, G., Wedel, B.J., Trebak, M., Bird, G.St.J., and Putney, J.W., Jr., Expression level of TRPC3 channel determines its mechanism of activation, J. Biol. Chem. 278, 21649, 2003. 61. Liu, X., Singh, B.B., and Ambudkar, I.S., TRPC1 is required for functional storeoperated Ca2+ channels. Role of acidic amino acid residues in the S5-S6 region, J. Biol. Chem. 278, 11337, 2003. 62. Brough, G.H., Wu, S., Ciofﬁ, D., Moore, T.M., Li, M., Dean, N., and Stevens, T., Contribution of endogenously expressed Trp1 to a Ca2+-selective, store-operated Ca2+ entry pathway, FASEB J. 15, 1727, 2001. 63. Vandebrouck, C., Martin, D., Colson-Van Schoor, M., Debaix, H., and Gailly, P., Involvement of TRPC in the abnormal calcium inﬂux observed in dystrophic (mdx) mouse skeletal muscle ﬁbers, J Cell Biol. 158, 1089, 2002. 64. Baldi, C., Vazquez, G., Calvo, J.C., and Boland, R., TRPC3-like protein is involved in the capacitative cation entry induced by 1alpha,25-dihydroxy-vitamin D3 in ROS 17/2.8 osteoblastic cells, J. Cell Biochem. 90, 197, 2003. 65. Wang, X., Pluznick, J.L., Wei, P., Padanilam, B.J., and Sansom, S.C., TRPC4 forms store-operated Ca2+ channels in mouse mesangial cells, Am. J. Physiol. Cell Physiol.287, C357, 2004. 66. Philipp, S., Trost, C., Warnat, J., Rautmann, J., Himmerkus, N., Schroth, G., Kretz, O., Nastainczyk, W., Cavalié, A., Hoth, M., and Flockerzi, V., Trp4 (CCE1) protein is part of native calcium release-activated Ca2+-like channels in adrenal cells, J. Biol. Chem. 275, 23965, 2000. 67. Freichel, M., Suh, S.H., Pfeifer, A., Schweig, U., Trost, C., Weißgerber, P., Biel, M., Philipp, S., Freise, D., Droogmans, G., Hofmann, F., Flockerzi, V., and Nilius, B., Lack of an endothelial store-operated Ca2+ current impairs agonist-dependent vasorelaxation in TRP4-/- mice, Nature Cell Biol. 3, 121, 2001. 68. Mori, Y., Wakamori, M., Miyakawa, T., Hermosura, M., Hara, Y., Nishida, M., Hirose, K., Mizushima, A., Kurosaki, M., Mori, E., Gotoh, K., Okada, T., Fleig, A., Penner, R., Iino, M., and Kurosaki, T., Transient receptor potential 1 regulates capacitative Ca2+ entry and Ca2+ release from endoplasmic reticulum in B lymphocytes, J. Exp. Med. 195, 673, 2002. 69. Zagranichnaya, T.K., Wu, X., and Villereal, M.L., Endogenous TRPC1, TRPC3, and TRPC7 proteins combine to form native store-operated channels in HEK-293 cells, J. Biol. Chem. 2005. 70. Tesfai, Y., Brereton, H.M., and Barritt, G.J., A diacylglycerol-activated Ca2+ channel in PC12 cells (an adrenal chromafﬁn cell line) correlates with expression of the TRP-6 (transient receptor potential) protein, Biochem. J. 358, 717, 2001. 71. Jung, S., Strotmann, R., Schultz, G., and Plant, T.D., TRPC6 is a candidate channel involved in receptor-stimulated cation currents in A7r5 smooth muscle cells, Am. J. Physiol. 282, C347, 2002. 72. Levchenko, A., Bruck, J., and Sternberg, P.W., Scaffold proteins may biphasically affect the levels of mitogen-activated protein kinase signaling and reduce its threshold properties, Proc. Nat. Acad. Sci. USA 97, 5818, 2000. 73. Burack, W.R. and Shaw, A.S., Signal transduction: hanging on a scaffold, Current Opinion in Cell Biology 12, 211, 2000. 74. Hoth, M. and Penner, R., Depletion of intracellular calcium stores activates a calcium current in mast cells, Nature 355, 353, 1992. 75. Hoth, M. and Penner, R., Calcium release-activated calcium current in rat mast cells, J. Physiol. (Lond. ) 465, 359, 1993.

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TRP Ion Channel Function in Sensory Transduction 76. Voets, T., Prenen, J., Fleig, A., Vennekens, R., Watanabe, H., Hoenderop, J.G.J., Bindels, R.J.M., Droogmans, G., Penner, R., and Nilius, B., CaT1 and the calcium release-activated calcium channel manifest distinct pore properties, J. Biol. Chem. 276, 47767, 2001. 77. Zhang, H., Inazu, M., Weir, B., Buchanan, M., and Daniel, E., Cyclopiazonic acid stimulates Ca2+ inﬂux through nonspeciﬁc cation channels in endothelial cells, Eur. J. Pharmacol. 251, 119, 1994. 78. Krause, E., Pfeiffer, F., Schmid, A., and Schulz, I., Depletion of intracellular calcium stores activates a calcium-conducting nonselective cation current in mouse pancreatic acinar cells, J. Biol. Chem. 271, 32523, 1996. 79. Wayman, C.P., Wallace, P., Gibson, A., and McFadzean, I., Correlation between storeoperated cation current and capacitative Ca2+ inﬂux in smooth muscle cells from mouse anococcygeus, Eur. J. Pharmacol. 376, 325, 1999. 80. Trepakova, E.S., Gericke, M., Hirakawa, Y., Weisbrod, R.M., Cohen, R.A., and Bolotina, V.M., Properties of a native cation channel activated by Ca2+ store depletion in vascular smooth muscle cells, J. Biol. Chem. 276, 7782, 2001. 81. McDaniel, S., Platoshyn, O., Wang, J., Yu, Y., Sweeney, M., Krick, S., Rubin, L.J., and Yuan, J.X.J., Capacitative Ca2+ entry in agonist-induced pulmonary vasoconstriction, Am. J. Physiol. 280, L870, 2001. 82. Albert, A.P. and Large, W.A., A Ca2+-permeable nonselective cation channel activated by depletion of internal Ca2+ stores in single rabbit portal vein myocytes, J. Physiol. (Lond. ) 538, 717, 2002. 83. Liu, X., Groschner, K., and Ambudkar, I.S., Distinct Ca(2+)-permeable cation currents are activated by internal Ca(2+)-store depletion in RBL-2H3 cells and human salivary gland cells, HSG and HSY, J. Membr. Biol. 200, 93, 2004. 84. Gusev, K., Glouchankova, L., Zubov, A., Kaznacheyeva, E., Wang, Z., Bezprozvanny, I., and Mozhayeva, G.N., The store-operated calcium entry pathways in human carcinoma A431 cells: functional properties and activation mechanisms, J. Gen. Physiol. 122, 81, 2003. 85. Philipp, S., Strauss, B., Hirnet, D., Wissenbach, U., Mery, L., Flockerzi, V., and Hoth, M., TRPC3 mediates T-cell receptor-dependent calcium entry in human Tlymphocytes, J. Biol. Chem. 278, 26629, 2003. 86. Hassock, S.R., Zhu, M.X., Trost, C., Flockerzi, V., and Authi, K.S., Expression and role of TRPC proteins in human platelets: evidence that TRPC6 forms the storeindependent calcium entry channel, Blood 100, 2801, 2002. 87. Inoue, R., Okada, T., Onoue, H., Hara, Y., Shimizu, S., Naitoh, S., Ito, Y., and Mori, Y., The transient receptor potential protein homologue TRP6 is the essential component of vascular α1-adrenoceptor-activated Ca2+-permeable cation channel, Circ. Res. 88, 325, 2001. 88. Gamberucci, A., Giurisato, E., Pizzo, P., Tassi, M., Giunti, R., McIntosh, D.P., and Benedetti, A., Diacylglycerol activates the inﬂux of extracellular cations in Tlymphocytes independently of intracellular calcium-store depletion and possibly involving endogenous TRP6 gene products, Biochem. J. 364, 245, 2002. 89. Albert, A.P. and Large, W.A., Synergism between inositol phosphates and diacylglycerol on native TRPC6-like channels in rabbit portal vein myocytes, J. Physiol. 552, 789, 2003. 90. Thebault, S., Zholos, A., Enﬁssi, A., Slomianny, C., Dewailly, E., Roudbaraki, M., Parys, J., and Prevarskaya, N., Receptor-operated Ca(2+) entry mediated by TRPC3/ TRPC6 proteins in rat prostate smooth muscle (PS1) cell line, J. Cell Physiol. 2005.

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91. Greka, A., Navarro, B., Oancea, E., Duggan, A., and Clapham, D.E., TRPC5 is a regulator of hippocampal neurite length and growth cone morphology, Nat. Neurosci. 6, 837, 2003. 92. Putney, J.W., Jr., Physiological mechanisms of TRPC activation, Pﬂugers Arch. 2005. 93. Vazquez, G., Wedel, B.J., Kawasaki, B.T., Bird, G.S., and Putney, J.W., Jr., Obligatory role of Src kinase in the signaling mechanism for TRPC3 cation channels, J. Biol. Chem. 279, 40521, 2004. 94. Trebak, M., Hempel, N., Wedel, B.J., Smyth, J.T., Bird, G.S., and Putney, J.W., Jr., Negative regulation of TRPC3 channels by protein kinase C–mediated phosphorylation of serine 712, Mol. Pharmacol. 67, 558, 2005. 95. Ahmmed, G.U., Mehta, D., Vogel, S., Holinstat, M., Paria, B.C., Tiruppathi, C., and Malik, A.B., Protein kinase C alpha phosphorylates the TRPC1 channel and regulates store-operated Ca2+ entry in endothelial cells, J Biol. Chem. 279, 20941, 2004.

3

TRPC2 and the Molecular Biology of Pheromone Detection in Mammals Emily R. Liman University of Southern California

Catherine Dulac Harvard University

CONTENTS Abstract ....................................................................................................................45 Introduction..............................................................................................................45 The Molecular Biology of Pheromone Sensing in Mammals................................46 TRPC2 Protein Structure and Binding Partners .....................................................48 TRPC2 Mechanism of Activation ...............................................................49 TRPC2 and the Evolution of Pheromone Sensing .....................................49 From VNO Activation to Behavioral Changes ...........................................50 Conclusion ...............................................................................................................51 References................................................................................................................51

ABSTRACT The expression of the TRPC2 channel in rodents appears largely restricted to neurons of the vomeronasal organ (VNO) and is detected at lower levels in few other tissues. The characteristics of TRPC2 expression, as well as the availability of the TRPC2 gene as a tool for genetic manipulation, has led to signiﬁcant advances in understanding the biology of the vomeronasal organ and pheromone detection in rodents and across evolution.

INTRODUCTION In order to ensure reproductive success, animals have evolved sensory and behavioral strategies to identify, respond to, and attract suitable mating partners. In many vertebrate and invertebrate species, chemical cues called pheromones carry the 45

46

TRP Ion Channel Function in Sensory Transduction

species- and gender-speciﬁc information required for mating. Detection of pheromones triggers the activation of likely hard-wired brain circuits, which in turn lead to stereotyped changes in the behavior and endocrine state of the animal. Molecular and physiological approaches have recently explored how the information provided by pheromones is detected in the nose and how the activation of deﬁned sets of pheromone receptors is translated into speciﬁc behavioral and physiological outputs.9 Molecular studies of chemosensory systems have made tremendous progress and opened new avenues of research with the discovery of receptor gene families and essential components of the signal transduction cascade in the main olfactory epithelium (MOE) and the vomeronasal organ (VNO).

THE MOLECULAR BIOLOGY OF PHEROMONE SENSING IN MAMMALS Surgical removal of olfactory and vomeronasal structures has traditionally assigned the role of the main olfactory system to the sense of smell, resulting in the detection of a large variety of volatile odorants,1,5 while the vomeronasal system is thought to carry most of the detection of gender- and species-speciﬁc cues involved in the control of mating and aggressive behavior.11 In the nasal cavity, volatile chemical cues interact with olfactory receptors (ORs) expressed in the MOE, while nonvolatile signals carried in the nasal mucus are actively pumped into the lumen of the VNO, where they can activate vomeronasal receptors (VRs) (Figure 3.1A and B). VNO neurons send their axonal projections to a distinct area of the dorsal telecephalon, the accessory olfactory bulb (AOB). Unlike mitral cells of the main olfactory bulb (MOB) that mainly innervate cortical areas, AOB mitral cells bypass the cerebral cortex and project directly to the medial amygdala, which, in turn, innervates nuclei of the hypothalamus involved in aggression and reproduction. Sensory neurons located in the apical layer of the VNO neuroepithelium each express a single member of the V1R family of VNO receptors and project to the anterior AOB, while neurons of the basal half of the neuroepithelium express receptors of the V2R gene family and send axons to the posterior AOB (Figure 3.1B) (reviewed in reference 9). The two murine VR gene families include 150–200 functional genes each and although both belong to the G-protein coupled receptor (GPCR) gene superfamily, they are phylogenetically unrelated to each other and to the ORs. Recent studies indicate thatV1R-expressing neurons respond to low molecular weight organic molecules11,18,44 while V2R-expressing cells respond to peptides.15,17 The V2Rs require the speciﬁc interaction with MHC Class Ib molecules M10s in order to be expressed in vitro and in vivo.24 Neurons in the main olfactory system and vomeronasal system utilize distinct signaling components to transduce chemical stimuli. The main components of the olfactory transduction cascade—Golf, ACIII, and OCNC1—are not expressed in vomeronasal sensory neurons,2 suggesting the existence of a different pathway for sensory transduction in the VNO. Because of similarity with phototransduction in the Drosophila eye30 and chemosensation in C. elegans,8 it was suggested that a TRP homologue might be involved in vertebrate pheromone signal transduction.20 Indeed, TRP2/TRPC2, a cation channel belonging to the transient receptor potential

TRPC2 and the Molecular Biology of Pheromone Detection in Mammals

47

FIGURE 3.1 (Color ﬁgure follows p. 234.) (A) The vomeronasal system: pheromone signals are detected by neurons of the vomeronasal organ (VNO) in the ventral nasal septum. VNO axons project to the accessory olfactory bulb (AOB), which in turn connects to nuclei of the vomeronasal amygdala in the limbic system. (B) Key signaling components of the vomeronasal organ: VNO neurons express one member of the two families of vomeronasal receptors V1Rs and V2Rs. V2Rs form a functional complex with the MHC class Ib molecules M10s. The activities of V1R- and V2R-expressing neurons require the expression of the TRPC2 channel. (C) Localization of rTRPC2 to the sensory microvilli of VNO neurons: anti-TRPC2 immunoﬂuorescence on sections of rat VNO demonstrates the localization of TRPC2 to the luminal surface of the VNO neurepithelium where pheromone-induced signaling is likely to occur. (Modiﬁed from reference 20.) (D) TRPC2 in primate evolution. Mutations that disrupt the opening of reading frame (ORF) of the TRPC2 protein were mapped to the time in primate evolution at which they ﬁrst occurred. The ﬁrst mutations (6 and 9) occurred in the ancestors of Old World monkeys and apes, at the same time that these animals developed trichromatic vision. Howler monkeys have independently evolved trichromatic vision but have an intact TRPC2 gene, indicating the trichromatic vision is not in itself sufﬁcient to replace pheromone signaling.37 (Modiﬁed from reference 21.) (E) The role of VNO signaling in gender discrimination and aggression: behavioral analysis of TRPC2-/- males reveals the essential role of the VNO in controlling the sex speciﬁcity of reproductive behavior, and in male-male aggression. Other sensory cues, most likely olfactory, are essential to trigger mating behavior in mice.

48

TRP Ion Channel Function in Sensory Transduction

(TRP) family of ion channels, appears highly expressed in VNO neurons and speciﬁcally localized to the microvilli, the proposed site of pheromone transduction (Figure 3.1C), suggesting a direct role in the VNO signaling cascade.

TRPC2 PROTEIN STRUCTURE AND BINDING PARTNERS TRPC2 was ﬁrst identiﬁed as a pseudogene in the database of human-expressed sequence tags (ESTs), and its presence in other genomes was conﬁrmed through sequencing of partial clones from mice and cows.38,39,43 A full-length sequence of rodent TRPC2 was subsequently identiﬁed simultaneously by two groups of researchers.20,36 The TRPC2 cDNA isolated from the rat vomeronasal organ (rTRPC2) by Liman et al. encodes a protein of 885 amino acids,20 and its mouse orthologue (mTRPC2β) encodes a protein of 890 amino acids that is 96 percent similar.12 Two long forms of mouse TRPC2 were identiﬁed by Vannier and colleagues (clones 14 and 17 or mTRPC2a and mTRPC2b)36; the predicted proteins of 1,172 and 1,072 amino acids, respectively, differ from the protein expressed in the VNO in that they contain extended N-termini. Subsequently a shorter variant (886 amino acids) of mTRPC2 (clone α)12 (see also reference 40) and three variants of mTRPC2 that encode proteins consisting of only portions of the extended N-terminal region from clones 14 and 17 were reported.7,40 The variant expressed in the VNO has been conﬁrmed by isolating the full cDNA from multiple cDNA libraries and through comparison of the size of the encoded protein with that of the native protein;20 moreover, the functionality of TRPC2 in the VNO has been clearly demonstrated (see below). For these reasons, unless otherwise noted, “TRPC2” will refer to the VNO splice variant. The possibility that other splice variants exist and have distinct functional roles is intriguing and merits additional study. The highest expression of TRPC2 is in sensory neurons of the vomeronasal organ, where it represents as much as 1/10,000 of the mRNA (Figure 3.1C).20 In these cells, the TRPC2 protein is remarkably localized to the sensory microvilli,20,28 actin-based structures that are specialized for the detection of chemical signals. Outside the vomeronasal organ, TRPC2 expression is low;12,20 it has been detected in the main olfactory epithelium,20 testes, heart, brain, liver, spleen, and erythrocytes.12,39 Notably, these tissues do not express the VNO form of TRPC2 (mTRPC2β), suggesting that they represent priming from alternate promoters. In sperm, TRPC2 protein has been detected in the acrosome region, suggesting a possible role in fertilization.14 The TRPC2 protein is structurally related to other TRP family members and is predicted to have, like these proteins and more distantly related voltage-gated K+ channels, six transmembrane domains and the ability to tetramerize. TRPC2 does not heteromultimerize with other TRPC channel subunits13 (but see reference 6), and thus native channels may either form homotetramers or may multimerize with more distantly related channel subunits. The predicted cytoplasmic N-terminus contains ankryn repeat domains, and the cytoplasmic C-terminus contains a coiled coil domain, both of which might mediate protein–protein interactions.20,36 Interaction between TRPC2 and the IP3 receptor has been demonstrated,4,35 and a conserved motif in the C-terminus of TRPC2 has been identiﬁed that binds the IP3 receptor

TRPC2 and the Molecular Biology of Pheromone Detection in Mammals

49

and calmodulin.34 Calmodulin has been shown to interact with the N-terminus of the long form of TRPC2.40 A novel protein named enkurin was identiﬁed from a yeast-two hybrid screen with the N-terminus of TRPC2, and the function of this protein is as yet unknown.33 Given the difﬁculty in studying the functional properties of TRPC2 (see below), the signiﬁcance of these protein interactions and of the domain structure of TRPC2 is not known.

TRPC2 MECHANISM

OF

ACTIVATION

Understanding the mechanism of activation of TRPC2 is critical to understanding its role in pheromone detection and other physiological processes. Despite the importance of this problem, it has remained refractory to study and there is presently no single agreed-upon mechanism for its activation. In heterologous cells, mTRPC2 (splice variants A and B) was reported to be activated by depletion of Ca2+ stores by thapsigargin and to function as a capacitative Ca2+ entry channel.10,35,36 Other experiments, however, showed that in heterologous cells TRPC2 (splice variants A, α, and β) is largely trapped in the endoplasmic reticulum, impeding study of its functional properties.12 (ER Liman, D Liu and J Appler, unpublished) An alternative is to study native TRPC2 channels. In sperm cells, thapsigargin induces a rise in Ca2+ that can be partially blocked by an antibody against an extracellular domain of TRPC2, suggesting that in these cells TRPC2 may be store operated.14 In sensory neurons from the VNO, TRPC2 is unlikely to be activated by depletion of Ca2+ stores, because the channel is localized in sensory microvilli at a considerable distance from Ca2+ stores.20,28 In these cells, TRPC2 may be gated by diacylglycerol, a conclusion based on the observation that VNO sensory neurons from TRPC2 knockout animals are missing a DAG-gated conductance found in wild-type cells.25 It is possible that there are different modes of activation of TRPC2, depending on the splice variant or the cell type in which it is expressed. Resolution of this controversy will require the successful expression of TRPC2 in heterologous cell types or its reintroduction into native cells.

TRPC2

AND THE

EVOLUTION

OF

PHEROMONE SENSING

The essential function and nearly exclusive expression of TRPC2 in the vomeronasal organ have made it an excellent marker to study changes in VNO function during evolution. In ﬁsh, which do not have a structurally distinct VNO, TRPC2 is expressed in the olfactory epithelium in a population of apical microvillar cells that also express VRs, and it is not expressed in the basal ciliated cells that express ORs.31 The microvillar cells appear specialized for detecting amino acids23 and send segregated projections to the lateral portion of the olfactory bulb.31 It is thus likely that the VNO arose by segregation of the microvillar cells from the ciliated cells, possibly as a response to terrestrial life. The main olfactory epithelium is well suited for detecting airborne chemicals that enter the nasal cavity during the respiratory cycle, whereas the VNO is better suited for detecting nonvolatile chemicals whose delivery is based on the presence of coinciding sensory and neuroendocrine signals.26 Whether humans have a functional VNO has been difﬁcult to determine using histological or functional techniques, and therefore it has been the subject of

50

TRP Ion Channel Function in Sensory Transduction

intense debate.29 The observation that the TRPC2 gene is a pseudogene in humans,21,38 as are most of the vomeronasal receptors of the V1R family,16 is strong evidence that the human VNO is vestigial. When did humans lose a functional VNO? A comprehensive analysis of the TRPC2 gene in extant primates has revealed that TRPC2 is a pseudogene in all Old World monkeys and apes, but not in New World monkeys (Figure 3.1D).21,42 Analysis of nucleotide substitutions during the predicted evolution of the TRPC2 gene shows that it changed through relaxed selective pressure in the ancestors of New World monkeys and apes, coincident with the development of trichromatic vision through duplication of the green opsin gene.21 It is likely that, at that time in evolution, visual signaling replaced the use of pheromones in communicating social and reproductive status. This conclusion is consistent with strong sexual dimorphism seen in Old World monkeys and primates.

FROM VNO ACTIVATION

TO

BEHAVIORAL CHANGES

The surgical ablation of the VNO or the AOB in rodents has been shown to impair mating and territorial defense of the animal, thus suggesting a critical role in gender and social recognition.11 However, clear differences in the degree of behavioral defects were observed in different species and, to some extent, in various reports of the same experiment. This can be easily explained by the behavioral variability existing between different species and strains and by the inherent difﬁculty of the surgical procedure performed on a sensory organ prone to regeneration. Genetic ablation of TRPC2 in mice provided a new experimental system to assess the requirement of TRPC2 function in VNO signaling and to directly investigate the repertoire of VNO-mediated sensory responses and behaviors.19,32 First, it appears that the TRPC2 deﬁciency dramatically impairs the sensory activation of VNO neurons by urine pheromones, thus conﬁrming the critical role of TRPC2 in the VNO signal transduction cascade. In addition, the absence of pheromone detection mediated by VNO signaling has striking behavioral consequences. TRPC2-/- male mice appear unable to recognize the sexual identity of their conspeciﬁcs: they fail to display the pheromone-evoked aggression toward male intruders that is normally seen in wild-type males and, remarkably, they display courtship and mounting behavior indiscriminately toward both males and females. These data contradict the established notion that VNO activity is required for the initiation of male-female mating behavior in mice and suggest instead a critical role in ensuring sex discrimination (Figure 3.1E). Defects in maternal aggression, male territory marking, and recognition of social dominance also appear impaired in the TRPC2-/- mouse line. The ability of TRPC2-/- males to mate, although indiscriminately with conspeciﬁcs of both sexes, emphasizes the essential role played by other sensory modalities in the control of reproductive behavior. Indeed, the VNO does not appear to be the sole detector of pheromonal cues. Recent evidence demonstrated that areas of the hypothalamus controlling reproduction and fertility receive inputs from the MOE.3,41 Likewise, odorant detection processed in the MOB is involved in the attraction of females toward males,22 and olfactory cues recently emerged as essential stimuli for reproductive behavior.27,41

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51

CONCLUSION The discovery of TRPC2 expression in the vomeronasal organ has led to major breakthroughs in understanding the molecular and cellular processes of pheromone signal transduction. Moreover, it has provided an essential tool to investigate the role of pheromone and vomeronasal signaling in organisms and across evolution. However, the most mechanistic aspects of TRPC2 function remain to be uncovered. Progress in this direction should provide novel insights into the translation of chemosensory detection into electrical signals and lead to a deeper understanding of the sensory coding of pheromones.

REFERENCES 1. L. Belluscio, G.H. Gold, A. Nemes, and R. Axel, Mice deﬁcient in G(olf) are anosmic, Neuron 20 (1998) 69–81. 2. A. Berghard, L.B. Buck, and E.R. Liman, Evidence for distinct signaling mechanisms in two mammalian olfactory sense organs, Proc. Natl. Acad. Sci. USA 93 (1996) 2365–2369. 3. U. Boehm, Z. Zou, and L.B. Buck, Feedback loops link odor and pheromone signaling with reproduction, Cell 123 (2005) 683–695. 4. J.H. Brann, J.C. Dennis, E.E. Morrison, and D.A. Fadool, Type-speciﬁc inositol 1,4,5-trisphosphate receptor localization in the vomeronasal organ and its interaction with a transient receptor potential channel, TRPC2, J. Neurochem. 83 (2002) 1452–1460. 5. L.J. Brunet, G.H. Gold, and J. Ngai, General anosmia caused by a targeted disruption of the mouse olfactory cyclic nucleotide-gated cation channel, Neuron 17 (1996) 681–693. 6. X. Chu, Q. Tong, J.Y. Cheung, J. Wozney, K. Conrad, V. Mazack, W. Zhang, R. Stahl, D.L. Barber, and B.A. Miller, Interaction of TRPC2 and TRPC6 in erythropoietin modulation of calcium inﬂux, J. Biol. Chem. 279 (2004) 10514–10522. 7. X. Chu, Q. Tong, J. Wozney, W. Zhang, J.Y. Cheung, K. Conrad, V. Mazack, R. Stahl, D.L. Barber, and B.A. Miller, Identiﬁcation of an N-terminal TRPC2 splice variant which inhibits calcium inﬂux, Cell Calcium 37 (2005) 173–182. 8. H.A. Colbert, T.L. Smith, and C.I. Bargmann, OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans, J. Neurosci. 17 (1997) 8259–8269. 9. C. Dulac and A.T. Torello, Molecular detection of pheromone signals in mammals: from genes to behaviour, Nat. Rev. Neurosci. 4 (2003) 551–562. 10. P. Gailly and M. Colson-Van Schoor, Involvement of trp-2 protein in store-operated inﬂux of calcium in ﬁbroblasts, Cell Calcium 30 (2001) 157–165. 11. M. Halpern, The organization and function of the vomeronasal system, Annual review of Neuroscience 10 (1987) 325–362. 12. T. Hofmann, M. Schaefer, G. Schultz, and T. Gudermann, Cloning, expression and subcellular localization of two novel splice variants of mouse transient receptor potential channel 2, Biochem. J. 351 (2000) 115–122. 13. T. Hofmann, M. Schaefer, G. Schultz, and T. Gudermann, Subunit composition of mammalian transient receptor potential channels in living cells, Proc. Natl. Acad. Sci. USA 99 (2002) 7461–7466.

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TRP Ion Channel Function in Sensory Transduction 14. M.K. Jungnickel, H. Marrero, L. Birnbaumer, J.R. Lemos, and H.M. Florman, Trp2 regulates entry of Ca2+ into mouse sperm triggered by egg ZP3, Nat. Cell Biol. 3 (2001) 499–502. 15. H. Kimoto, S. Haga, K. Sato, and K. Touhara, Sex-speciﬁc peptides from exocrine glands stimulate mouse vomeronasal sensory neurons, Nature 437 (2005) 898–901. 16. H. Kouros-Mehr, S. Pintchovski, J. Melnyk, Y.J. Chen, C. Friedman, B. Trask, and H. Shizuya, Identiﬁcation of nonfunctional human VNO receptor genes provides evidence for vestigiality of the human VNO, Chem. Senses 26 (2001) 1167–1174. 17. T. Leinders-Zufall, P. Brennan, P. Widmayer, P. Chandramani S., A. Maul-Pavicic, M. Jager, X.H. Li, H. Breer, F. Zufall, and T. Boehm, MHC class I peptides as chemosensory signals in the vomeronasal organ, Science 306 (2004) 1033–1037. 18. T. Leinders-Zufall, A.P. Lane, A.C. Puche, W. Ma, M.V. Novotny, M.T. Shipley, and F. Zufall, Ultrasensitive pheromone detection by mammalian vomeronasal neurons, Nature 405 (2000) 792–796. 19. B.G. Leypold, C.R. Yu, T. Leinders-Zufall, M.M. Kim, F. Zufall, and R. Axel, Altered sexual and social behaviors in trp2 mutant mice, Proc. Natl. Acad. Sci. USA 99 (2002) 6376–6381. 20. E.R. Liman, D.P. Corey, and C. Dulac, TRP2: a candidate transduction channel for mammalian pheromone sensory signaling, Proc. Natl. Acad. Sci. USA 96 (1999) 5791–5796. 21. E.R. Liman and H. Innan, Relaxed selective pressure on an essential component of pheromone transduction in primate evolution, Proc. Natl. Acad. Sci. USA 100 (2003) 3328–3332. 22. Y. Lin da, S.Z. Zhang, E. Block, and L.C. Katz, Encoding social signals in the mouse main olfactory bulb, Nature 434 (2005) 470–477. 23. D.L. Lipschitz and W.C. Michel, Amino acid odorants stimulate microvillar sensory neurons, Chem. Senses 27 (2002) 277–286. 24. J. Loconto, F. Papes, E. Chang, L. Stowers, E.P. Jones, T. Takada, A. Kumanovics, K. Fischer Lindahl, and C. Dulac, Functional expression of murine V2R pheromone receptors involves selective association with the M10 and M1 families of MHC class Ib molecules, Cell 112 (2003) 607–618. 25. P. Lucas, K. Ukhanov, T. Leinders-Zufall, and F. Zufall, A diacylglycerol-gated cation channel in vomeronasal neuron dendrites is impaired in TRPC2 mutant mice: mechanism of pheromone transduction, Neuron 40 (2003) 551–561. 26. M. Luo, M.S. Fee, and L.C. Katz, Encoding pheromonal signals in the accessory olfactory bulb of behaving mice, Science 299 (2003) 1196–1201. 27. V.S. Mandiyan, J.K. Coats, and N.M. Shah, Deﬁcits in sexual and aggressive behaviors in Cnga2 mutant mice, Nat. Neurosci. 8 (2005) 1660–1662. 28. B.P. Menco, V.M. Carr, P.I. Ezeh, E.R. Liman, and M.P. Yankova, Ultrastructural localization of G-proteins and the channel protein TRP2 to microvilli of rat vomeronasal receptor cells, J. Comp. Neurol. 438 (2001) 468–489. 29. M. Meredith, Human vomeronasal organ function: a critical review of best and worst cases, Chem. Senses 26 (2001) 433–445. 30. B.A. Niemeyer, E. Suzuki, K. Scott, K. Jalink, and C.S. Zuker, The Drosophila lightactivated conductance is composed of the two channels TRP and TRPL, Cell 85 (1996) 651–659. 31. Y. Sato, N. Miyasaka, and Y. Yoshihara, Mutually exclusive glomerular innervation by two distinct types of olfactory sensory neurons revealed in transgenic zebraﬁsh, J. Neurosci. 25 (2005) 4889–4897.

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32. L. Stowers, T.E. Holy, M. Meister, C. Dulac, and G. Koentges, Loss of sex discrimination and male-male aggression in mice deﬁcient for TRP2, Science 295 (2002) 1493–1500. 33. K.A. Sutton, M.K. Jungnickel, Y. Wang, K. Cullen, S. Lambert, and H.M. Florman, Enkurin is a novel calmodulin and TRPC channel-binding protein in sperm, Dev. Biol. 274 (2004) 426–435. 34. J. Tang, Y. Lin, Z. Zhang, S. Tikunova, L. Birnbaumer, and M.X. Zhu, Identiﬁcation of common binding sites for calmodulin and inositol 1,4,5-trisphosphate receptors on the carboxyl termini of trp channels, J. Biol. Chem. 276 (2001) 21303–21310. 35. Q. Tong, X. Chu, J.Y. Cheung, K. Conrad, R. Stahl, D.L. Barber, G. Mignery, and B.A. Miller, Erythropoietin-modulated calcium inﬂux through TRPC2 is mediated by phospholipase Cgamma and IP3R, Am. J. Physiol. Cell Physiol. 287 (2004) C1667–1678. 36. B. Vannier, M. Peyton, G. Boulay, D. Brown, N. Qin, M. Jiang, X. Zhu, and L. Birnbaumer, Mouse trp2, the homologue of the human trpc2 pseudogene, encodes mTrp2, a store depletion–activated capacitative Ca2+ entry channel, Proc. Natl. Acad. Sci. USA 96 (1999) 2060–2064. 37. D.M. Webb, L. Cortes-Ortiz, and J. Zhang, Genetic evidence for the coexistence of pheromone perception and full trichromatic vision in howler monkeys, Mol. Biol. Evol. 21 (2004) 697–704. 38. P.D. Wes, J. Chevesich, A. Jeromin, C. Rosenberg, G. Stetten, and C. Montell, TRPC1, a human homolog of a Drosophila store-operated channel, Proc. Natl. Acad. Sci. USA 92 (1995) 9652–9656. 39. U. Wissenbach, G. Schroth, S. Philipp, and V. Flockerzi, Structure and mRNA expression of a bovine trp homologue related to mammalian trp2 transcripts, FEBS Lett. 429 (1998) 61–66. 40. E. Yildirim, A. Dietrich, and L. Birnbaumer, The mouse C-type transient receptor potential 2 (TRPC2) channel: alternative splicing and calmodulin binding to its N-terminus, Proc. Natl. Acad. Sci. USA 100 (2003) 2220–2225. 41. H. Yoon, L.W. Enquist, and C. Dulac, Olfactory inputs to hypothalamic neurons controlling reproduction and fertility, Cell 123 (2005) 669–682. 42. J. Zhang and D.M. Webb, Evolutionary deterioration of the vomeronasal pheromone transduction pathway in catarrhine primates, Proc. Natl. Acad. Sci. USA 100 (2003) 8337–8341. 43. X. Zhu, M. Jiang, M. Peyton, G. Boulay, R. Hurst, E. Stefani, and L. Birnbaumer, trp, a novel mammalian gene family essential for agonist-activated capacitative Ca2+ entry, Cell 85 (1996) 661–671. 44. K. Del Punta, T. Leinders-Zufall, I. Rodriguez, D. Jukam, C.J. Wysocki, S. Ogawa, F. Zufall, and P. Mombaerts, Deﬁcient pheromone responses in mice lacking a cluster of vomeronasal receptor genes, Nature 419 (2002) 70–74.

4

TRP Channels and Axon Pathﬁnding Kai Cui and Xiao-bing Yuan Chinese Academy of Sciences

CONTENTS Abstract ....................................................................................................................55 Overview of Nerve Pathﬁnding...............................................................................55 Calcium Signal in Growth Cone Turning ...............................................................56 TRPCs’ Function in Nerve Guidance......................................................................57 TRPC5 Regulates Nerve Growth ................................................................57 TRPC Contributes to Nerve Pathﬁnding.....................................................58 TRPC Gating Mechanisms by Guidance Factors .......................................59 How TRPC Channels Contribute to Guidance Signaling...........................59 Possible Roles of TRP Channels in Neuronal Migration.......................................61 Calcium Signal in Neuronal Migration.......................................................61 Mechanosensation in Migrating Neurons?..................................................61 Thermosensation for Migration? .................................................................63 Conclusion ...............................................................................................................63 Acknowledgments....................................................................................................63 References................................................................................................................63

ABSTRACT Transient receptor potential channels (TRP) are a large family of cationic channels that are permeable to Ca2+, allowing these channels to participate in a wide range of physiological processes that require Ca2+ signaling. In recent years, TRP channels have been found to play a role in many processes in the nervous system, such as the transduction of sensory stimulation,1 neuronal cell death,2 proliferation and differentiation of neural progenitor cells,3,4 nerve growth,5 and synaptic transmission.6–9 One interesting recent discovery shows that TRPC (canonical) channels are involved in the signal transduction of axon guidance during brain development.

OVERVIEW OF NERVE PATHFINDING As the most complicated organ in animals, the brain is composed of highly ordered cell architecture and precisely wired neural circuits, which are essential for the 55

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sophisticated functions of the brain. The development of functional neuronal circuits requires the ﬁnely tuned growth of nerve ﬁbers toward their targets, a process called nerve pathﬁnding.10 Both the axon and dendrites of neurons have the pathﬁnding process during development, and some mechanisms were shared by these two different neuronal processes.11,12 Therefore, in this chapter when we talk about the nerve growth or nerve guidance, we do not discuss these two processes separately without special mention. At the tip of growing nerves there is a highly dynamic structure called a growth cone, which bears motile ﬁlopodia and lamellipodia, exploring the environment and leading the nerve growth toward its favorite direction. During brain development, it is generally believed that concentration gradients of the guidance cues exist in the tissue.13,14 When the growth cone is exposed to a concentration gradient of a chemoattractant or chemorepellent, it responds by turning toward or away from the source of the guidance cue, one of the basic behaviors of growth cones during nerve pathﬁnding. During this response, receptors on the growth cone’s surface can read the concentration gradient and transduce the direction signal into the cytosol, initiating a series of downstream events, including the rearrangement of the actin and microtubule cytoskeletons inside the cell,15 the changing of adhesion level at different sides of the growth cone,16 polarized endocytosis and exocytosis of vesicles containing membrane proteins, synthesis and insertion of some new proteins for nerve pathﬁnding,17–19 and degradation of some proteins to control the local protein level.20 Transduction of some long-range signals back to the soma and other neurites in the same cell may be also required for the neuron to coordinate the response of different neurites in response to the stimuli.21–23 Eventually, the growth cone will turn and grow stably in the new direction.

CALCIUM SIGNAL IN GROWTH CONE TURNING With regard to the complicated signal transduction process during this growth cone turning triggered by guidance cues, one fundamental question is what kind of intracellular signal can be activated to mediate the directional information after the growth cone reads the concentration gradient of extracellular guidance cues. The most attractive candidate for this mediator of the growth cone guidance signal is the intracellular Ca2+ ([Ca2+]i),24 which regulates the motility of the growth cone and the response of growth cones to many extracellular factors.25–27 Using an in vitro growth cone guidance model,28 previous studies showed that many extracellular factors that can guide the nerve growth, including netrin-1, slit, brain-derived neurotrophic factor (BDNF), myelin-associated glycoprotein (MAG), SDF-1, and several neurotranmitters,22,29–33 can elevate the [Ca2+]i in the growth cone. Preventing the [Ca2+]i elevation in growth cones by removing the extracellular Ca2+ (for some factors), by depleting the intracellular Ca2+ store, or by blocking the Ca2+ release from internal stores, inhibits the growth cone turning triggered by a concentration gradient of guidance cues.24 A gradient of [Ca2+]i at the growth cone is also sufﬁcient to drive the nerve growth cone turning because an artiﬁcially generated concentration gradient of [Ca2+]i, either by local photolysis of caged compound of Ca2+ or by an extracellular gradient of ryanodine (a drug that opens the internal store of Ca2+), can trigger the growth cone to extend toward the higher [Ca2+]i side.29,34 Moreover, Ca2+

TRP Channels and Axon Pathﬁnding

57

has been shown to regulate a wide range of intracellular signaling molecules involved in nerve growth, including those microtubule- and actin-associated proteins, motor proteins, kinases, phosphotases, and proteases.24 Based on these facts, a hypothesis is widely accepted that a concentration gradient of extracellular guidance factors can be translated into a concentration gradient of [Ca2+]i across the growth cone, which then drives the preferential growth of the nerve toward the higher [Ca2+]i side. Under such a hypothesis, it is of great interest to know how the Ca2+ signal is triggered by guidance factors. Elevation of [Ca2+]i can result from Ca2+ inﬂux through voltage-gated Ca2+ channels in plasma membrane in response to membrane depolarization or from the opening of ligand-gated Ca2+ channels, for example, ionotropic glutamate receptors. Internal release of Ca2+ triggered by the messenger molecule inositol-1,4,5-trisphosphate (InsP3) or by Ca2+ itself may also contribute to elevation of [Ca2+]i.24 Another important pathway that can lead to the elevation of [Ca2+]i is the Ca2+ inﬂux through the voltage-independent channels including the TRP channels, which may be gated by both receptor activation and store-operated mechanisms.1,35 Previous studies showed that both the Ca2+ inﬂux through membrane channels and the Ca2+ release from internal stores are required for the growth cone turning triggered by a group of guidance molecules including netrin, BDNF, and MAG.13,29,30,36 In these studies, it was shown that netrin triggers the opening of the L-type Ca2+ channel, an ion channel that is opened upon membrane depolarization, and Ca2+ inﬂux through this voltage-gated Ca2+ channel is required for the attractive growth cone turning triggered by netrin.29,37 Observations from a lot of other researchers also showed that during early development spontaneous neuronal activity is required for the proper growth and nerve pathﬁnding through elevating the [Ca2+]i.30,38,39 One unavoidable question raised by these studies is how could membrane potential be elevated during pathﬁnding of these immature neurons? This puzzle obtained an attractive answer by the exciting recent discovery that TRPC channels, whose openings are not voltage dependent, play essential roles in the axon pathﬁnding.

TRPCS’ FUNCTION IN NERVE GUIDANCE TRPC5 REGULATES NERVE GROWTH The ﬁrst clue comes from the neurite extension study carried out in cultured hippocampal neurons.5 In order to address the function of TRPC5 channels that are expressed at high levels in the hippocampus, the dominant negative mutant proteins of TRPC5 were expressed in cultured hippocampal neurons. One interesting ﬁnding is that overexpression of wide-type TRPC5 in these neurons dramatically reduced the neurite extension rate. On the contrary, expression of dominant negative TRPC5 resulted in increased ﬁlopodia length and an elevated neurite extension rate. Consistently, a substantial amount of TRPC5 protein can be detected at the growth cone probed with a speciﬁc anti-TRPC5 antibody. The researchers concluded from these observations that spontaneous opening of TRPC5 may mediate the inﬂux of Ca2+ from extracellular media, which may be adverse for the neurite extension. Since the

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TRPC5 channel regulates the growth cone morphology and the neurite extension rate, it is highly possible that these Ca2+-permeable channels may be involved in the growth cone pathﬁnding, which is the neurite extension controlled by extracellular factors.

TRPC CONTRIBUTES

TO

NERVE PATHFINDING

New ﬁndings veriﬁed this prediction. Three independent studies showed that TRPC channels may be involved in the attractive growth cone turning triggered by the chemoattractant netrin or BDNF in cultured cerebellar granule cells of rat and embryonic spinal neurons of Xenopus.40–42 In these studies, growth cone turning assay was used to test the response of the growth cone to an extracellular gradient of guidance factors.16,28 A micropipette containing guidance molecules at a very high dosage was used to generate a concentration gradient of the molecule through repetitive pufﬁng, and the gradient source (the pipette tip) was positioned 100 μm in front of the growth cone at an angle of 45 degrees with respect to the original direction of the nerve extension. The growth cone can sense this concentration gradient of guidance molecules and migrate toward or away from the gradient, depending on what guidance molecules are tested. Interestingly, all these studies showed that chemoattraction triggered by BDNF and netrin was blocked by inhibiting TRP channels with a general inhibitor SKF-96365 or by speciﬁc knocking down of the expression of endogenous TRPC protein with small interference RNA (siRNA) or morpholino-antisense RNA. Meanwhile, the [Ca2+]i elevation triggered by netrin and BDNF at the growth cone was also dramatically reduced after perturbing the TRPC channels, as revealed by Ca2+ imaging or patch-clamp recording at the growth cone. This suggests that in response to the stimulation of BDNF and netrin, TRPC channels may mediate the Ca2+ inﬂux, which is required for triggering the growth cone turning. Moreover, the cation inﬂux through TRP channels may elevate the level of membrane potential, leading to the activation of a voltagedependent sodium channel and calcium channels, which further depolarize membrane potential and allow more Ca2+ inﬂux. These studies also showed that the role of TRPC channels in growth cone turning is speciﬁc. In rat granule neurons, TRPC3 and 6 are only required for BDNF-triggered growth cone attraction but not for either growth cone attraction triggered by glutamate or repulsion triggered by SDF-1. In the Xenopus spinal neurons, the chemoattractant netrin and BDNF as well as the repellent MAG engage Xenopus TRP-1 (xTRPC1) to guide the growth cone, while another guidance molecule (semaphorin) doesn’t use it.42 The requirement of TRPC channels in the axon guidance in vivo was further demonstrated by Shim et al.42 Apart from a similar observation using the growth cone turning assay, they looked at the growth of the commissural axons in the spinal cord after injection of the morphalino RNA of the xTRPC1 in one cell at the twocell stage of the embryo to silence the expression of xTRPC1 proteins during development. They found that knocking down the expression of xTRPC1 dramatically deferred the midline cross of these commissural axons and caused some axons to go astray, suggesting that the xTRPC1 channel can mediate the axon guidance signal in vivo. Although currently no TRPC knockout mice have been reported to

TRP Channels and Axon Pathﬁnding

59

have nerve pathﬁnding defects, it may be due to the compensation of protein function by other TRP family members when a single TRPC gene is knocked out. It is possible that defects in nerve pathﬁnding may be seen in mice of compound knockout of several family members. Not only chemoattraction requires TRPC channels, it was also shown that the repulsive growth cone turning triggered by MAG was also eliminated by blocking the xTRPC channels.42 Because MAG is one of the molecules that prevents axon regeneration after the injury of central nervous systems in adult animals, the engagement of the TRPC channel in MAG signaling suggests that these ion channels will be candidate drug targets to enhance nerve regeneration.

TRPC GATING MECHANISMS

BY

GUIDANCE FACTORS

One study explored the opening mechanism of the TRPC channel in the growth cone by the guidance molecule BDNF.40 Similar to the PLC dependency of many other TRP channels, the researchers found that both the growth cone attraction and the elevation of [Ca2+]i triggered by BDNF were blocked by inhibition of PLC signaling with the speciﬁc inhibitor U73122. In contrast, inhibition of PKC activity did not have any effect. Furthermore, blocking the InsP3 signaling with xestospongin C or 2APB, which perturb the InsP3 receptor at the internal store of Ca2+, also blocked the growth cone turning and [Ca2+]i elevation, suggesting that Ca2+ release from internal stores may be required for the opening of TRPC channels in the plasma membrane. Overall, these observations support such a model where the binding of BDNF with its receptor TrkB activates PLC-γ, which generates IP343 (see Figure 4.1). The subsequent opening of IP3 induces the Ca2+ release from internal stores. This Ca2+ elevation can further activate the TRPC channels in the plasma membrane and allow more Ca2+ influx. In such a model, TRPC channels in the growth cone membrane should be activated through some Ca2+-dependent mechanisms (Figure 4.1). This was supported by the recent report that the activation of TRPC5 by agonists can be regulated by calmodulin and [Ca2+]i.44 Because many extracellular factors that have been shown to regulate nerve growth can activate PLC activity through receptor tyrosine kinase or G-protein-coupled receptors that are linked to the signaling of PLC-γ and PLC-β, respectively,45,46 PLC-dependent TRP channel activation may also mediate Ca2+ signaling for other guidance factors during nerve pathﬁnding. However, how the activation of guidance receptors is coupled to the opening of TRPC channels, and whether there is direct association of guidance receptors with the TRPC protein like the binding of TrkB with TRPC3,47 remains to be clarified.

HOW TRPC CHANNELS CONTRIBUTE

TO

GUIDANCE SIGNALING

Among various TRPC proteins, TRPC3, TRPC6, and TRPC7 belong to one subfamily35,48 and can form functional heteromeric or homomeric ion channels.49,50 In cultured hippocampal neurons, expression of DN-TRPC5 elevates neurite extension, whereas overexpression of wild-type TRPC5 inhibits nerve growth.5 However, downregulation of TRPC3/6 did not signiﬁcantly affect neurite growth in cerebellar granule cells.40 This differential effect on nerve growth may owe to longer mean channel open

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FIGURE 4.1 TRPC channels in nerve guidance. An extracellular gradient of guidance cue (from left upper corner) triggers local [Ca2+]i elevation, establishing a [Ca2+]i gradient across the growth cone. The graded calcium signal causes imbalanced polymerization/stablization of actin and microtubule elements and polarized transport of membrane vesicles (circle) to drive the growth cone extending toward the higher Ca2+ side. Calcium elevation is achieved by the binding of the guidance cue (e.g., BDNF or netrin) with its receptor (TrkB or DCC) on the surface of the growth cone, leading to the activation of phospholipase-γ (PLC-γ), which catalyzes the production of IP3. The activation of IP3R at the endoplasmic reticulum triggers Ca2+ release from internal stores. This process is coupled to the mobilization of TRPC channels in the plasma membrane through unknown mechanisms, resulting in Ca2+ inﬂux. The membrane depolarization triggered by the opening of TRPC channels may activate VDCCs, which bring in additional Ca2+ inﬂux. TRP channels at the soma and other parts of the migrating neurons are supposed to act as sensors for temperature and mechanical forces.

time of TRPC5-containing channels than TRPC3/6 channels. That allows much larger Ca2+ inﬂux,51–53 leading to adverse effects to neurite growth, whereas a modest level of Ca2+ inﬂux through a TRPC3/6 channel is sufﬁcient to trigger attractive growth cone turning. By allowing different patterns of Ca2+ inﬂux, diverse TRPC channels may thus carry out distinct regulatory functions at the growth cone. Previous studies have divided the guidance molecules in two groups depending on the requirement of extracellular Ca2+ and the modulation properties by cyclic AMP (cAMP) or cyclic GMP (cGMP) during the growth cone turning triggered by these factors.13 Type I factors require the extracellular Ca2+ to trigger the growth cone turning, and cAMP level can switch the turning response of the growth cone, with a higher level of cAMP leading to chemoattraction and a lower level to repulsion. In contrast, the growth cone turning triggered by type II guidance molecules is independent of extracellular Ca2+ and is switched by intracellular cGMP level. Intracellular calcium signaling (with either type I or II factors) is required for

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61

the growth cone turning response.13 It is possible that TRP channels participate in the growth cone guidance triggered by type I factors, by which the Ca2+ release from internal stores is not sufﬁcient to trigger the growth cone turning. For the type II factors, however, the Ca2+ release from internal stores upon receptor activation may be enough to drive the turning responses. Meanwhile, the opening of ryanodine channels at the internal stores also contributes to the Ca2+ signaling in netrin-triggered chemoattraction.29 For the type I guidance factors, TRPC channels may not necessarily be the sole membrane ion channels engaged in the nerve pathﬁnding. The opening of these cationic channels allows the depolarization of membrane potential in response to the stimulation of guidance molecules. In consequence, voltage-gated cationic channels will open and trigger the further depolarization of the membrane potential, accompanied with inﬂux of Ca2+ through voltage-dependent calcium channels.41 Calcium-triggered Ca2+ release from internal stores may also be triggered as a consequence of [Ca2+]i elevation. Calcium from all these sources may contribute to nerve growth and nerve pathﬁnding in some neuronal tissues; however, TRPC activation is the trigger of signal ampliﬁcation in response to type I guidance factors.

POSSIBLE ROLES OF TRP CHANNELS IN NEURONAL MIGRATION CALCIUM SIGNAL

IN

NEURONAL MIGRATION

Apart from nerve pathﬁnding, other processes in neural development may also be regulated by TRP channels. One such developmental process is neuronal migration. Unlike nerve growth, neuronal migration is the translocation of the whole cell body including the leading process and the soma. However, some signaling processes may be shared by both nerve pathﬁnding and neuronal migration. One example is that a series of guidance factors—slit, netrin, ephrins, BDNF, SDF—have been shown to be used for both nerve guidance and the guidance of neuronal migration.54 Moreover, the Ca2+ signal has been shown to regulate the speed and direction of neuronal migration.55–58 One example is the neuronal migration in postnatal cerebellum tissue, in which a high frequency of Ca2+ waves is correlated with the high migrating speed of the young granule cells,55 and the end of the Ca2+ wave is linked to the cease of the migration.58 NMDA receptor activity and N-type Ca2+ channels are required to maintain the normal migration of these neurons.59 Therefore, it is highly possible that TRPC channels that participate in the nerve guidance signaling, and whose openings allow Ca2+ inﬂux into the cytosol, may also play a role in controlling the motility and the guidance of neuronal migration. Determining which TRPC proteins are involved in neuronal migration deserves further study.

MECHANOSENSATION

IN

MIGRATING NEURONS?

During its migration, stress and tension are generated toward the neuron because it has to attach and squeeze in surrounding tissue, which is ﬁlled with other cells and an extracellular matrix. The same is true for the exploring growth cone, although

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the size of the growth cone is relatively smaller. These mechanical forces and their change over time may reﬂect current motility and position of the migrating cell. It is likely that cells sense and use this information to ﬁne-tune their pace and direction of migration, such as changing their shapes in order to select a way of stronger adhesion and lower resistance to their movements. Consistent with this possibility, it was frequently observed that neurons change their shapes dramatically during migration including becoming highly elongated.60,61 Therefore, it is reasonable to hypothesize that mechanical sensing may play an important role in neuronal migration, helping the neuron to actively reorganize the cytoskeleton, to change its shape, or even to reduce and recover its volume in order to ﬁght its way in tissues. In agreement with this prediction, mechanically stimulated currents were recorded in the growth cones of cultured snail neurons and rat DRG neurons.62,63 More interestingly, ﬁlopodia, which play an essential role in exploring the surrounding tissue, were the most sensitive to mechanical stimulation. Mechanical tension can be an important regulator of nerve growth because neurite outgrowth can be initiated de novo by experimental application of tension to the cell margin of neurons in culture prior to spontaneous neurite outgrowth.64–66 The “towed” neurites had a normal appearance, showed rapid saltatory movements of internal organelles, and were capable of sustained growth on the substratum. Electron microscopy of bundles of neurites produced in this way from explanted dorsal root ganglia showed an ultrastructure typical of cultured neurites, with abundant longitudinally aligned microtubules and neuroﬁlaments.64,67 In cultured hippocampal neurons, applied mechanical tension could stimulate undifferentiated minor processes to become axons and could induce a second axon in an already polarized neuron.68 In a cultured ﬁbroblast, it was reported that stretch-activated Ca2+ entry sustains cell migration and mediates active and passive responses to mechanical signals.69 In this study, application of gadolinium, an inhibitor of stretch-activated ion channels, or removal of extracellular-free Ca2+, caused inhibition of traction forces. Gadolinium treatment also inhibited cell migration.69 Immunoﬂuorescence microscopy indicated that gadolinium caused a dramatic decrease in vinculin and phosphotyrosine concentrations at focal adhesions, suggesting that stretch-activated Ca2+ entry regulates the organization of focal adhesions and the output of mechanical forces. Likewise, mechanosensory channels may be involved in the migration of neurons during development. So far, the molecular nature for these mechanical sensors at the migrating growth cone and soma remains unknown. Some researchers reported that the N-type calcium channel on the cell membrane is mechanosensitive.70 Interestingly, N-type calcium channels have been reported to play a role in directed migration of immature neurons as indicated by the reductive effect of the channel’s inhibitor on the migration of cultured cerebellar granule cells.71 In contrast, inhibitors of L- and T-type calcium channels, as well as those of sodium and potassium channels, had no effect on the rate of granule cell migration.71 Because gadolinium, a drug previously used to inhibit stretch-activated ion channels, can also block or activate TRP channels at different concentrations,72 it is unclear whether this drug reduced the migration of ﬁbroblasts69 through inhibiting TRP channels. The recent discovery that TRP channels play essential roles in mechanical sensing in sensory neurons1,73 raised the

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63

possibility that these TRP channels may play a role in sensing the adhesion and compactness of the surrounding tissue of migrating neurons. Whether the mechanosensitive TRP channels in sensory neurons are also expressed in migrating neurons during development and whether they really serve as the mechanical sensor and inﬂuence neuronal migration will be fascinating subjects for further investigation.

THERMOSENSATION

FOR

MIGRATION?

Imaging studies in brain slice cultures showed that neuronal migration is highly sensitive to temperature.59,60,74 Considering that TRP channels play essential roles in heat and cold sensing1,35 and that some of these TRP channels are also expressed in the brain,35,75 one reasonable supposition is that these channels also sense temperature when the neurons migrate in a normal tissue environment, allowing Ca2+ inﬂux and maintaining a suitable level of Ca2+ in the cytosol to keep the motility of the cells. Exploring the expression patterns of TRP family members in the brain at different developmental stages will help to determine which TRP proteins may be actively involved in speciﬁc developmental processes.

CONCLUSION Evidence is emerging that Ca2+-permeable TRP channels regulate nerve growth and pathﬁnding. The Ca2+ permeability and various gating mechanisms suggest that TRP channels may be widely involved in other developmental processes including neuronal migration, dendrite arborization, spine generation, and synapse formation. How TRP channels are opened by various physiological stimuli during development remains unclear. The next several years will witness the identiﬁcation of more functions played by TRP channels in various developmental processes. The further clariﬁcation of the function of TRP channels and the underlying mechanisms in neural development will dramatically increase our understanding of how brain circuits are connected.

ACKNOWLEDGMENTS We would like to thank Dr. Yi-zheng Wang, Ning Li, and Chen-bing Guan for comments on the manuscript.

REFERENCES 1. Moran, M.M., Xu, H., and Clapham, D.E. TRP ion channels in the nervous system. Curr. Opin. Neurobiol. 14, 362–69 (2004). 2. Aarts, M.M. and Tymianski, M. TRPMs and neuronal cell death. Pﬂugers Arch. (2005). 3. Pla, A.F. et al. Canonical transient receptor potential 1 plays a role in basic ﬁbroblast growth factor (bFGF)/FGF receptor-1-induced Ca2+ entry and embryonic rat neural stem cell proliferation. J. Neurosci. 25, 2687–701 (2005).

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TRP Ion Channel Function in Sensory Transduction 4. Wu, X., Zagranichnaya, T.K., Gurda, G.T., Eves, E.M., and Villereal, M.L. A TRPC1/ TRPC3-mediated increase in store-operated calcium entry is required for differentiation of H19-7 hippocampal neuronal cells. J. Biol. Chem. 279, 43392–402 (2004). 5. Greka, A., Navarro, B., Oancea, E., Duggan, A., and Clapham, D.E. TRPC5 is a regulator of hippocampal neurite length and growth cone morphology. Nat. Neurosci. 6, 837–45 (2003). 6. Gee, C.E., Benquet, P., and Gerber, U. Group I metabotropic glutamate receptors activate a calcium-sensitive transient receptor potential-like conductance in rat hippocampus. J. Physiol. 546, 655–64 (2003). 7. Tozzi, A. et al. Involvement of transient receptor potential-like channels in responses to mGluR-I activation in midbrain dopamine neurons. Eur. J. Neurosci. 18, 2133–145 (2003). 8. Bengtson, C.P., Tozzi, A., Bernardi, G., and Mercuri, N.B. Transient receptor potential-like channels mediate metabotropic glutamate receptor EPSCs in rat dopamine neurones. J. Physiol. 555, 323–30 (2004). 9. Munsch, T., Freichel, M., Flockerzi, V., and Pape, H.C. Contribution of transient receptor potential channels to the control of GABA release from dendrites. Proc. Natl. Acad. Sci. USA 100, 16065–70 (2003). 10. Tessier-Lavigne, M. and Goodman, C.S. The molecular biology of axon guidance. Science 274, 1123–33 (1996). 11. Polleux, F., Morrow, T., and Ghosh, A. Semaphorin 3A is a chemoattractant for cortical apical dendrites. Nature 404, 567–73 (2000). 12. Kim, S. and Chiba, A. Dendritic guidance. Trends Neurosci. 27, 194–202 (2004). 13. Song, H.J. and Poo, M.M. Signal transduction underlying growth cone guidance by diffusible factors. Curr. Opin. Neurobiol. 9, 355–63 (1999). 14. Tessier-Lavigne, M. Axon guidance by molecular gradients. Curr. Opin. Neurobiol. 2, 60–65 (1992). 15. Song, H. and Poo, M. The cell biology of neuronal navigation. Nat. Cell Biol. 3, E81–88 (2001). 16. Gundersen, R.W. and Barrett, J.N. Characterization of the turning response of dorsal root neurites toward nerve growth factor. J. Cell Biol. 87, 546–54 (1980). 17. Ming, G.L. et al. Adaptation in the chemotactic guidance of nerve growth cones. Nature 417, 411–18 (2002). 18. Campbell, D.S. and Holt, C.E. Chemotropic responses of retinal growth cones mediated by rapid local protein synthesis and degradation. Neuron 32, 1013–26 (2001). 19. Wu, K.Y. et al. Local translation of RhoA regulates growth cone collapse. Nature 436, 1020–24 (2005). 20. Campbell, D.S. and Holt, C.E. Apoptotic pathway and MAPKs differentially regulate chemotropic responses of retinal growth cones. Neuron 37, 939–52 (2003). 21. Jellies, J., Loer, C.M., and Kristan, W.B., Jr. Morphological changes in leech Retzius neurons after target contact during embryogenesis. J. Neurosci. 7, 2618–29 (1987). 22. Zheng, J.Q., Zheng, Z., and Poo, M. Long-range signaling in growing neurons after local elevation of cyclic AMP-dependent activity. J. Cell Biol. 127, 1693–701 (1994). 23. Davenport, R.W., Thies, E., and Cohen, M.L. Neuronal growth cone collapse triggers lateral extensions along trailing axons. Nat. Neurosci. 2, 254–59 (1999). 24. Henley, J. and Poo, M.M. Guiding neuronal growth cones using Ca2+ signals. Trends Cell Biol. 14, 320–30 (2004). 25. Kater, S.B. and Mills, L.R. Regulation of growth cone behavior by calcium. J. Neurosci. 11, 891–99 (1991).

TRP Channels and Axon Pathﬁnding

65

26. Kater, S.B., Davenport, R.W., and Guthrie, P.B. Filopodia as detectors of environmental cues: signal integration through changes in growth cone calcium levels. Prog. Brain Res. 102, 49–60 (1994). 27. Gomez, T.M., Robles, E., Poo, M., and Spitzer, N.C. Filopodial calcium transients promote substrate-dependent growth cone turning. Science 291, 1983–87 (2001). 28. Lohof, A.M., Quillan, M., Dan, Y., and Poo, M.M. Asymmetric modulation of cytosolic cAMP activity induces growth cone turning. J. Neurosci. 12, 1253–61 (1992). 29. Hong, K., Nishiyama, M., Henley, J., Tessier-Lavigne, M., and Poo, M. Calcium signalling in the guidance of nerve growth by netrin-1. Nature 403, 93–98 (2000). 30. Ming, G., Henley, J., Tessier-Lavigne, M., Song, H., and Poo, M. Electrical activity modulates growth cone guidance by diffusible factors. Neuron 29, 441–52 (2001). 31. Xu, H.T. et al. Calcium signaling in chemorepellant slit2-dependent regulation of neuronal migration. Proc. Natl. Acad. Sci. USA 101, 4296–301 (2004). 32. Jin, M. et al. Ca2+-dependent regulation of rho GTPases triggers turning of nerve growth cones. J. Neurosci. 25, 2338–47 (2005). 33. Xiang, Y. et al. Nerve growth cone guidance mediated by G-protein-coupled receptors. Nat. Neurosci. 5, 843–48 (2002). 34. Zheng, J.Q. Turning of nerve growth cones induced by localized increases in intracellular calcium ions. Nature 403, 89–93 (2000). 35. Clapham, D.E., Runnels, L.W., and Strübing, C. The TRP ion channel family. Nat. Rev. Neurosci. 2, 387–96 (2001). 36. Ming, G.L. et al. cAMP-dependent growth cone guidance by netrin-1. Neuron 19, 1225–35 (1997). 37. Nishiyama, M. et al. Cyclic AMP/GMP-dependent modulation of Ca2+ channels sets the polarity of nerve growth-cone turning. Nature 423, 990–95 (2003). 38. McFarlane, S. and Pollock, N.S. A role for voltage-gated potassium channels in the outgrowth of retinal axons in the developing visual system. J. Neurosci. 20, 1020–29 (2000). 39. Pollock, N.S. et al. Voltage-gated potassium channels regulate the response of retinal growth cones to axon extension and guidance cues. Eur. J. Neurosci. 22, 569–78 (2005). 40. Li, Y. et al. Essential role of TRPC channels in the guidance of nerve growth cones by brain-derived neurotrophic factor. Nature 434, 894–98 (2005). 41. Wang, G.X. and Poo, M.M. Requirement of TRPC channels in netrin-1-induced chemotropic turning of nerve growth cones. Nature 434, 898–904 (2005). 42. Shim, S. et al. XTRPC1-dependent chemotropic guidance of neuronal growth cones. Nat. Neurosci. 8, 730–35 (2005). 43. Huang, E.J. and Reichardt, L.F. Trk receptors: roles in neuronal signal transduction. Annu. Rev. Biochem. 72, 609–42 (2003). 44. Ordaz, B. et al. Calmodulin and calcium interplay in the modulation of TRPC5 channel activity. Identiﬁcation of a novel C-terminal domain for calcium/calmodulinmediated facilitation. J. Biol. Chem. 280, 30788–96 (2005). 45. Montell, C. PLC ﬁlls a GAP in G-protein-coupled signalling. Nat. Cell Biol. 2, E82–83 (2000). 46. Rebecchi, M.J. and Pentyala, S.N. Structure, function, and control of phosphoinositidespeciﬁc phospholipase C. Physiol. Rev. 80, 1291–335 (2000). 47. Li, H.S., Xu, X.Z., and Montell, C. Activation of a TRPC3-dependent cation current through the neurotrophin BDNF. Neuron 24, 261–73 (1999).

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TRP Ion Channel Function in Sensory Transduction 48. Trebak, M., Vazquez, G., Bird, G.S., and Putney, J.W., Jr. The TRPC3/6/7 subfamily of cation channels. Cell Calcium 33, 451–61 (2003). 49. Hofmann, T., Schaefer, M., Schultz, G., and Gudermann, T. Subunit composition of mammalian transient receptor potential channels in living cells. Proc. Natl. Acad. Sci. USA 99, 7461–66 (2002). 50. Goel, M., Sinkins, W.G., and Schilling, W.P. Selective association of TRPC channel subunits in rat brain synaptosomes. J. Biol. Chem. 277, 48303–10 (2002). 51. Yamada, H. et al. Spontaneous single-channel activity of neuronal TRP5 channel recombinantly expressed in HEK293 cells. Neurosci. Lett. 285, 111–14 (2000). 52. Jung, S. et al. Lanthanides potentiate TRPC5 currents by an action at extracellular sites close to the pore mouth. J. Biol. Chem. 278, 3562–71 (2003). 53. Zitt, C. et al. Expression of TRPC3 in Chinese hamster ovary cells results in calciumactivated cation currents not related to store depletion. J. Cell Biol. 138, 1333–41 (1997). 54. Guan, K.L. and Rao, Y. Signalling mechanisms mediating neuronal responses to guidance cues. Nat. Rev. Neurosci. 4, 941–56 (2003). 55. Komuro, H. and Rakic, P. Intracellular Ca2+ ﬂuctuations modulate the rate of neuronal migration. Neuron 17, 275–85 (1996). 56. Kumada, T. and Komuro, H. Completion of neuronal migration regulated by loss of Ca(2+) transients. Proc. Natl. Acad. Sci. USA 101, 8479–84 (2004). 57. Komuro, H. and Rakic, P. Orchestration of neuronal migration by activity of ion channels, neurotransmitter receptors, and intracellular Ca2+ ﬂuctuations. J. Neurobiol. 37, 110–30 (1998). 58. Komuro, H. and Kumada, T. Ca2+ transients control CNS neuronal migration. Cell Calcium 37, 387–93 (2005). 59. Rakic, P. and Komuro, H. The role of receptor/channel activity in neuronal cell migration. J. Neurobiol. 26, 299–315 (1995). 60. Komuro, H. and Rakic, P. Dynamics of granule cell migration: a confocal microscopic study in acute cerebellar slice preparations. J. Neurosci. 15, 1110–20 (1995). 61. Komuro, H. and Rakic, P. Distinct modes of neuronal migration in different domains of developing cerebellar cortex. J. Neurosci. 18, 1478–90 (1998). 62. Sigurdson, W.J. and Morris, C.E. Stretch-activated ion channels in growth cones of snail neurons. J. Neurosci. 9, 2801–8 (1989). 63. Imai, K., Tatsumi, H., and Katayama, Y. Mechanosensitive chloride channels on the growth cones of cultured rat dorsal root ganglion neurons. Neuroscience 97, 347–55 (2000). 64. Bray, D. Axonal growth in response to experimentally applied mechanical tension. Dev. Biol. 102, 379–89 (1984). 65. Chada, S., Lamoureux, P., Buxbaum, R.E., and Heidemann, S.R. Cytomechanics of neurite outgrowth from chick brain neurons. J. Cell Sci. 110 (Pt. 10), 1179–86 (1997). 66. Zheng, J. et al. Tensile regulation of axonal elongation and initiation. J. Neurosci. 11, 1117–25 (1991). 67. Zheng, J., Buxbaum, R.E., and Heidemann, S.R. Investigation of microtubule assembly and organization accompanying tension-induced neurite initiation. J. Cell Sci. 104 (Pt. 4), 1239–50 (1993). 68. Lamoureux, P., Ruthel, G., Buxbaum, R.E., and Heidemann, S.R. Mechanical tension can specify axonal fate in hippocampal neurons. J. Cell Biol. 159, 499–508 (2002). 69. Munevar, S., Wang, Y.L., and Dembo, M. Regulation of mechanical interactions between ﬁbroblasts and the substratum by stretch-activated Ca2+ entry. J. Cell Sci. 117, 85–92 (2004).

TRP Channels and Axon Pathﬁnding

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70. Calabrese, B., Tabarean, I.V., Juranka, P., and Morris, C.E. Mechanosensitivity of N-type calcium channel currents. Biophys. J. 83, 2560–74 (2002). 71. Komuro, H. and Rakic, P. Selective role of N-type calcium channels in neuronal migration. Science 257, 806–9 (1992). 72. Tousova, K., Vyklicky, L., Susankova, K., Benedikt, J., and Vlachova, V. Gadolinium activates and sensitizes the vanilloid receptor TRPV1 through the external protonation sites. Mol. Cell Neurosci. 30, 207–17 (2005). 73. Lin, S.Y. and Corey, D.P. TRP channels in mechanosensation. Curr. Opin. Neurobiol. 15, 350–57 (2005). 74. Komuro, H., Yacubova, E., and Rakic, P. Mode and tempo of tangential cell migration in the cerebellar external granular layer. J. Neurosci. 21, 527–40 (2001). 75. Guatteo, E. et al. Temperature sensitivity of dopaminergic neurons of the substantia nigra pars compacta: involvement of TRP channels. J. Neurophysiol. (2005).

5

TRPV1 Receptors and Signal Transduction Tamara Rosenbaum Universidad Nacional Autónoma de México

Sidney A. Simon Duke University

CONTENTS Introduction..............................................................................................................69 TRPV1 Activators....................................................................................................72 Activation by Capsaicin and Other TRPV1 Agonists.................................72 TRPV1 Activation by Protons.....................................................................73 Noxious Heat Promotes TRPV1 Activation................................................74 Modulation of TRPV1 Activity by Cellular Components ......................................75 Phosphorylation by PKA.............................................................................76 Phosphorylation by PKC .............................................................................76 Phosphorylation by CaMKII and Binding and Modulation of TRPV1 Activity by CaM ........................................................................76 Modulation by Lipids ..................................................................................77 Effects of Reducing Agents on TRPV1 Activity ........................................77 Acknowledgments....................................................................................................78 References................................................................................................................78

INTRODUCTION The perception of pain throughout the body arises when neural signals originating from the terminals of nociceptors are propagated to second-order neurons in the spinal cord or brainstem, whereupon they are transmitted to speciﬁc higher order brain areas (Price, 2000). Recent studies have begun to elucidate some of the molecular mechanisms underlying the transduction of noxious stimuli. Many stimuli have been found to activate ion channels present on nociceptor terminals that act as molecular transducers to depolarize these neurons, thereby setting off nociceptive impulses along the pain pathways (Price, 2000; Costigan and Woolf, 2000). Among these ion channels are the members of the transient receptor potential (TRP) family. To date, the most studied member of the TRP family is the TRPV1 receptor. This is because it is the 69

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only one activated by capsaicin, the compound in chili pepper responsible for its “hot” taste; also, inhibiting TRPV1 has been shown to have therapeutic value (DiMarzo et al., 2002; Cortright and Szallasi, 2004). Although we will focus on the presence of these channels in nociceptors, we note that they have been identiﬁed in many other cell types and in various cortical and subcortical areas (Toth et al., 2005). The transient receptor potential vanilloid 1 (TRPV1) channel is predicted to have six transmembrane domains and a short, pore-forming hydrophobic stretch between the ﬁfth and sixth transmembrane domains (see Figure 5.1A). It is activated not only

FIGURE 5.1 (Color ﬁgure follows p. 234.) (A) A schematic diagram of a TRPV1 subunit in a bilayer. The subunit has six transmembrane domains (red) and a pore loop. The functional TRPV1 receptor is believed to form a tetramer. Residues involved in vanilloid binding are shown in orange. “A” indicates ankyrin repeats shown in yellow. Residues susceptible of phosphorylation are shown in green. Two calmodulin-binding regions in the N- and C-termini are indicated by “CaM.” Blue residues in the “P-loop” represent protonatable amino acids. Cysteine residues in the P-loop are susceptible of reduction and are indicated by the color purple. PIP2 is shown to bind to the region indicated in the C-terminus. The TRP box represents the TRP

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71

by the vanilloid capsaicin (Caterina et al., 1997), but also by noxious heat (>43°C) and low pH (Caterina et al., 1997; Tominaga et al., 1998), voltage (Gunthorpe et al., 2000; Piper et al., 1999), and various lipids (Julius and Basbaum, 2001; Caterina and Julius, 2001; Clapham, 2003; Cortright and Szallasi, 2004, Szallasi and Blumberg, 1999; Prescott and Julius, 2003; Jung et al., 2004; Bhave et al., 2003). In cells, TRPV1 is inactivated by its binding to PIP2 and is released from this block by PLC-mediated PIP2 hydrolysis (Prescott and Julius, 2003). Since its cloning in 1997, many amino acid regions within the TPRV1 protein have been shown to be involved in speciﬁc functions, such as capsaicin, proton, and heat activation; voltage dependence; permeability and ion selectivity; antagonist regions; desensitization; phosphorylation; modulation by lipids; and multimerization. In regard to its subunit composition, functional TRPV1 channels likely exist as homomeric or heteromeric complexes composed of four subunits that assemble to form functional cation-(including calcium) permeable pores (Clapham, 2003; Kedei et al., 2001; Kuzhikanathil et al., 2001). Moreover, like other ion channels, these channels have been shown to be associated with regulatory proteins (see Figure 5.1B and Kim et al., 2006). There are many signaling pathways that become activated (or inhibited) by the activation of TRPV1 (Farkas-Szallasi et al., 1995; Wood et al., 1988). Similar to many other channels, TRPV1 contains multiple phosphorylation sites in its amino acid sequence for protein kinase C (PKC) (Bhave et al., 2003; Dai et al., 2004; Premkumar et al., 2004), protein kinase A (PKA) (Bhave et al., 2002; De Petrocellis et al., 2001; Rathee et al., 2002) and Ca2+/calmodulin-dependent protein kinase II (CaMKII). The presence of multiple phosphorylation sites in TRPV1 implies possible regulatory actions by these kinases (Wood et al., 1988). Discussed later in this chapter are several lines of evidence that show that two types of lipids—endocannabinoids and eicosanoids that are products of lipoxygenase (LOX)—activate TRPV1 channels (Zygmunt et al., 1999; Hwang et al., 2000).

FIGURE 5.1 (Continued) domain. (Modiﬁed from Ferrer-Montiel et al., 2004 and from Tominaga and Tominaga, 2005.) (B) Role of metabolic pathways of anandamide and arachidonic acid in TRPV1 activation. The fatty acid amide hydrolase (FAAH) hydrolyzes anandamide (AEA) to produce arachidonic acid (AA) and ethanolamide. AA is oxygenated by lipoxygenase enzymes to produce TRPV1 agonists, 12- and 15-HPETE, 5-HETE and leukotriene B4 (LTB4) (Hwang et al., 2000). AEA, a substrate for lipoxygenase, yields equivalent HPETE ethanolamides (HPETEE) and HETE ethanolamides (HETEE) that are proposed to be TRPV1 agonists (Craib et al., 2001). The lipoxygenase products of anandamide act as potent inhibitors of FAAH (Maccarrone et al., 2000). PKC activates the TRPV1 receptor directly (Premkumar and Ahern, 2000; Olah et al., 2002), but it also sensitizes the receptor to other agonists (Vellani et al., 2001). AEA directly activates PKC (De Petrocellis et al., 1995; Premkumar and Ahern, 2000). CB1 receptor activation is coupled to PLC stimulation, PIP2 hydrolysis and release of the TRPV1 receptor from the inhibitory effect of this compound (see Hermann et al., 2003). AEA production occurs through phosphodiesterase-mediated cleavage of a phospholipid precursor, NAPE (N-arachidonoyl-phosphatidylethanolamine), in calcium-dependent fashion (DiMarzo et al., 1994). AEA is synthesized in response to TRPV1 receptor activation in cultured neurons (Ahluwalia et al., 2003). (Modiﬁed from Ross, 2003.)

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Because TRPV1 functions as a molecular integrator for multiple types of sensory input, in this chapter we will explore the molecular mechanisms underlying the activation and modulation of this channel.

TRPV1 ACTIVATORS Before proceeding with the identiﬁcation of regions on TRPV1 associated with the binding of activators, care must be taken that these are actually the binding sites as opposed to gating sites that will also lead to changes in the allosteric properties of the protein. This is important because, for example, capsaicin, even at concentrations that fail to activate a current, can sensitize TRPV1 receptors to protons and heat. Similarly, protons can sensitize TRPV1 receptors to capsaicin and heat. As discussed below, this likely occurs as a consequence of decreasing the free energy of one of the many closed states for these stimuli to be closer to one of the open states (Hui et al., 2003; Ryu et al., 2003). That is, TRPV1 is a multistate ion channel whose rate constants between the rate limiting closed and open states may depend on voltage, temperature, and a variety of agonists.

ACTIVATION

BY

CAPSAICIN

AND

OTHER TRPV1 AGONISTS

TRPV1 receptors are activated by vanilloids like capsaicin (Spath and Darling, 1930; Thresh, 1846). At negative holding potentials, this activation results in the inﬂux of calcium and sodium, thereby depolarizing the cell. TRPV1 can be activated by capsaicin in isolated membrane patches that are devoid of the intracellular signaling machinery. Judging from results obtained from binding assays (Szallasi et al., 1993) and from electrophysiological recordings (Hui et al., 2003), there is good agreement that the binding of at least two capsaicin molecules is required for complete activation of this channel. There is good evidence that the capsaicin-binding site is intracellular on TRPV1 receptors. One form of evidence is that when added extracellularly, membraneimpermeant forms of capsaicin are inactive, but are active when applied intracellularly (Jung et al., 1999). In the rodent form of TRPV1, residues Arg114 and Glu761 in the intracellular N- and C-termini, respectively, have been identiﬁed as agonist recognition sites (Jung et al., 2002). By analyzing the avian form of TRPV1 that is not activated by capsaicin with murine forms that are, it was found that residues Tyr511 and Thr550 in the third and ﬁfth transmembrane domains are responsible for binding capsaicin to TRPV1 (Jordt and Julius, 2002; Gavva et al., 2004). In addition, three residues located at the transition between the second intracellular loop and the third transmembrane domain (Tyr511 and Ser512) and at the bottom of the ﬁfth transmembrane domain (Tyr550) have been proposed as sites where vanilloid agonists might interact with the TRPV1 channel (Jordt and Julius, 2002; Gavva et al., 2004). One of the best characterized of the less-pungent capsaicin analogues is olvanil. Olvanil, like capsaicin, has nociceptive and inﬂammatory properties (Dray and Dickenson, 1991), and it is about as efﬁcient as capsaicin in its ability to increase Ca2+ inﬂux in cultured rat DRGs (Walpole et al., 1993; Koplas et al., 1997) and induce currents in nociceptive neurons (Liu and Simon, 1997). One reason that it is

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73

a less-pungent analogue is that its rate of activation of TRPV1 is slower than that of capsaicin; while TRPV1 is being activated and the receptor potential is slowly depolarizing, voltage-dependent sodium and calcium channels will become inactivated to a greater extent. This in turn will decrease the probability of this neuron ﬁring action potentials (Liu et al., 1997). Another reason is that olvanil also activates CB1 receptors, which leads to anti-nociceptive responses. An important advance was the identiﬁcation of resiniferatoxin (RTX), a diterpene related to the phorbol esters, as a potent capsaicin analogue (Szallasi and Blumberg, 1989). This compound, which shares a vanillyl group with capsaicin, is a particularly strong irritant that was isolated from the latex of a Moroccan cactus-like plant Euphorbia resinifera (Hergenhahn et al., 1984; Appendino and Szallasi, 1997). In several assays, RTX is several thousandfold more potent than capsaicin (Szolcsanyi et al., 1990). Rat and human TRPV1 have been pharmacologically characterized, proving that RTX is indeed an agonist of TRPV1 in DRG neurons. The use of radio-labeled RTX for TRPV1 has allowed the detection of TRPV1 in peripheral tissues such as the urinary bladder (Szallasi et al., 1993; Acs et al., 1994), urethra (Parlani et al., 1993), nasal mucosa (Rinder et al., 1996), airways (Szallasi et al., 1993), and colon (Goso et al., 1993), as well as in several brain nuclei (Mezey et al., 2000). RTX binding requires the presence of a methionine in position 547 of the third transmembranal segment (Gavva et al., 2004); these residues are involved also in the binding of some endogenous agonists as well as of competitive antagonists (Jordt and Julius, 2002; Gavva et al., 2004; Phillips et al., 2004). In addition to capsaicin and RTX, several other natural and synthetic hydrophobic TRPV1 agonists have been identiﬁed (Rami and Gunthorpe, 2004; Krause et al., 2005; Cortright and Szallasi, 2004). Among the natural agonists found in the body are the arachidonic acid metabolites anandamide (which also activates CB1 and TRPV4 receptors) and 12-hydroxyeicosatetranenoic acid (12-HETE) (see Figure 5.1B). In addition, N-arachidonoyldopamine (NADA) activates TRPV1, but its biological function remains unknown. Piperine and zingerone, two pungent tasting compounds found in black pepper and ginger, respectively, have also been shown to activate TRPV1 receptors (Liu and Simon, 1996; McNamara et al., 2005). In summary, TRPV1 can be activated by a variety of molecules, and many more are certain to be identiﬁed.

TRPV1 ACTIVATION

BY

PROTONS

Pain sensation is augmented by the acidic extracellular pH during ischemia or inﬂammation. It has been shown that Aδ and C-ﬁber neurons transduce extracellular acid signals (hydrated protons) by means of at least two different classes of cationselective channels, TRPV1 (Caterina et al., 1997; Gunthorpe et al., 2002) and the acid-sensing ion channels (ASICs) (Bianchi and Driscoll, 2002; Kellenberger et al., 2002; Waldmann et al., 1999). For TRPV1 receptors, it has been found that protons only activate the channels when they are added from the extracellular solution, suggesting that the protonactive gate is on the extracellular part of the molecule (Jordt et al., 2000; Welch et al., 2000). Lowering the pH causes sigmoidal increases in the current. This current has a reversal potential close to zero millivolts (Bevan and Yeats, 1991; Liu and

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Simon, 2000), suggesting that, at neutral pH, protons do not carry much of the current in buffers containing physiological concentrations of sodium and calcium. Nevertheless, under speciﬁc experimental conditions, it has recently been suggested that TRPV1 is permeable to protons and that the open TRPV1 pore holds enough water molecules to form a continuous “water wire,” allowing “proton hopping” along adjacent free water molecules (Hellwig et al., 2004). Protons inﬂuence the cloned TRPV1 in a complex fashion. Lowering the pH sensitizes the responses to capsaicin (Tominaga et al., 1998; Ryu et al., 2003; Reeh and Kress, 2002). This is in accordance with earlier observations, where low pH potentiated responses to low concentrations of capsaicin in sensory neurons in cultures from rats (Petersen and LaMotte, 1993; Kress et al., 1996; Liu and Simon, 2000), rabbits (Martenson et al., 1994), or humans (Baumann et al., 1996). The presence of protons increases the afﬁnity of the receptor for capsaicin and changes the distribution of energy states for capsaicin activation that will bias them toward the open state, thereby affecting the mean open times (Ryu et al., 2003). Future work will test speciﬁc downstream effects of the TRPV1-mediated dual acidiﬁcation mechanism in nociceptive neurons, but it is clear that low pHs potentiate activation of TRPV1 by capsaicin in cell-free patches where the intracellular machinery of signaling is nonexistent.

NOXIOUS HEAT PROMOTES TRPV1 ACTIVATION The application of temperature ramps revealed that TRPV1 acts as a molecular thermometer. At a holding potential of –60 mV, the inward current abruptly increases at about 43°C (a temperature called the transition temperature). If kept at that temperature (or above) the channels will desensitize. Threshold temperature is reduced at a lower pH and also in the presence of capsaicin (Guenther et al., 1999; Tominaga et al., 1998). Heat-evoked TRPV1 currents exhibit properties similar to those of capsaicinevoked currents. Single-channel openings elicited by heat are observed in inside-out membrane patches excised from HEK293 cells expressing TRPV1. Similar to what happens for capsaicin or proton-induced activation of TRPV1, because it can be activated even in excised membrane patches, this channel seems to be itself a heat sensor. It is now known that several TRP family ion channels (TRPV1, TRPV2, TRPV3, TRPV4, TRPM8, TRPA1) are thermosensitive. This suggests that temperature sensor domains must be present in these channels (Patapoutian et al., 2003; Numazaki and Tominaga, 2004). It has been proposed (Vlachova et al., 2003) and now demonstrated (Brauchi et al., 2006) that the distal half of the TRPV1 C-terminus is reportedly involved in thermal sensitivity. Moreover, it has been shown that certain mutations and phosphorylation by PKA or PKC lead to the reduction of the threshold temperature for TRPV1 activation (Numazaki et al., 2002; Tominaga et al., 2001). Nevertheless, no TRPV1 mutation has been reported to markedly abrogate the response to heat, which suggests more global effects of heat on TRPV1. It is worth mentioning that any results obtained in the future through mutagenesis of the channel have to be carefully analyzed to exclude the possibility that these manipulations affect channel gating by heat in an allosteric fashion.

TRPV1 Receptors and Signal Transduction

75

To date, several TRPV1 splice variants have been identiﬁed. In rats, an N-terminal deletion splice variant of TRPV1, VR.5′sv, (Xue et al., 2001; Schumacher et al., 2000) lacks the majority of the N-terminus (amino acids 1-308 and 345-404) and does not form functional channels. More recently, two murine splice variants, mTRPV1α and mTRPV1β, were identiﬁed (Wang et al., 2004a). It was found that mTRPV1α (but not mTRPV1β) was activated by capsaicin and protons, suggesting that mTRPV1α is activated in a similar manner to other TRPV1 channels. In contrast, the mTRPV1β subunit, characterized by a ten amino acid deletion near the N-terminus, acts as a naturally occurring dominant negative regulator. The human hTRPV1b splice variant differs from the human TRPV1 in that it contains a sixty amino acid deletion within the N-terminus, and in that it forms functional channels that are activated by noxious heat (threshold, 47°C) but not activated by capsaicin or protons. It has been suggested that it may serve predominantly as a thermal receptor for nociceptive heat stimuli (Lu et al., 2005). Although TRPV1 is clearly ligand gated, it is also known to have voltagedependent gating properties (Gunthorpe et al., 2000). Moreover, a recent report has tightly linked temperature sensing in TRPV1 to voltage-dependent gating although this is somewhat controversial. Changes in temperature result in graded shifts in the voltage-dependent activation curve of TRPV1 (Voets et al., 2004). The hypothesis is that there are amino acids responsible for voltage dependence and that these amino acids are also involved in thermosensing (Branchi et al., 2006), even though the fourth TM domain of TRPV1 lacks the multiple positively charged residues typically present in voltage-gated channels (Voets et al., 2004).

MODULATION OF TRPV1 ACTIVITY BY CELLULAR COMPONENTS In addition, TRPV1’s response to heat can be modiﬁed by tyrosine kinases or Gprotein-coupled receptors. In these cases, the channel opens even at a normal body temperature (Vellani et al., 2001; Tominaga et al., 2001). Hence, one important aspect of TRPV1 regulation that has received extensive attention concerns the molecular and cellular mechanisms by which the inﬂammatory mediators in damaged tissues sensitize TRPV1 to its chemical and physical stimuli. Similar to what happens in other ion channels, TRPV1 binds and is modulated by molecules such as calcium calmodulin (CaM) (Numazaki et al., 2003; Rosenbaum et al., 2004). It can be phosphorylated by kinases including PKA (Vlachova et al., 2003; De Petrocellis et al., 2001; Rathee et al., 2002), PKC (Bhave et al., 2003; Premkumar et al., 2004; Tominaga et al., 2001; Varga et al., 2006), calcium calmodulin– dependent kinase II (CaMKII) (Jung et al., 2004), or Src kinase (Jin et al., 2004). Further stimulation of TRPV1 activity can be achieved by inﬂammatory agents such as bradykinin, serotonin, histamine, or prostaglandins, which stimulate TRPV1 either by protein kinase C–dependent pathways (Cesare et al., 1999; Premkumar and Ahern, 2000; Vellani et al., 2001); by releasing the channel from phosphatidylinositol 4,5-bisphosphate-dependent inhibition (Chuang et al., 2001; Prescott and Julius, 2003); by a protein kinase A–mediated recovery from inactivation (Bhave et al., 2002); or by formation of 12-hydroperoxyeicosatetraenoic acid (12-HPETE) (Shin et al., 2002) (see Figure 5.1B).

76

PHOSPHORYLATION

TRP Ion Channel Function in Sensory Transduction BY

PKA

Inﬂammatory mediators such as prostaglandins promote the activation of a PKAdependent pathway inﬂuencing capsaicin- or heat-mediated actions of TRPV1 in sensory neurons. Therefore, it seems that PKA plays a crucial role in the development of hyperalgesia and inﬂammation. Residues Ser 116 and Thr 370 in the amino terminus are phosphorylated by PKA and implicated in desensitization (Bhave et al., 2002; Mohapatra and Nau, 2003). PKA has been shown to phosphorylate residue Ser 116, regulating TRPV1 activity. Finally, residues Thr 144, Thr 370, and Ser 502 have also been implicated in sensitization of heat-evoked TRPV1 responses when phosphorylated by PKA (Rathee et al., 2002).

PHOSPHORYLATION

BY

PKC

Phosphorylation of TRPV1 by PKC is a downstream event from activation of Gq-coupled receptors by several inﬂammatory mediators such as ATP, bradykinin, prostaglandins, and trypsin or tryptase (Moriyama et al., 2005; Moriyama et al., 2003; Tominaga et al., 1998; Cortright and Szallasi, 2004). Phosphorylation of TRPV1 by PKC acts to potentiate capsaicin- or proton-evoked responses and reduces the temperature threshold for TRPV1 activation. Direct phosphorylation of TRPV1 by PKC has been demonstrated (Numazaki et al., 2002), and two target Ser residues (Ser 502 and Ser 800) have been identiﬁed (Bhave et al., 2003; Numazaki and Tominaga, 2004). When these two Ser residues are replaced with Ala, TRPV1 activity induced by capsaicin, protons, or heat is eliminated. These two serines are also implicated in potentiation of endovanilloid/endocannabinoid N-Arachidonoyldopamine (NADA)-induced TRPV1 activation (Premkumar et al., 2004), oleoylethanolamide (OEA)-induced TRPV1 activation (Ahern, 2003), and rephosphorylation of TRPV1 after desensitization in the presence of Ca2+ (Mandadi et al., 2004). While some investigators have provided support for the involvement of PKC (Cesare et al., 1999; Khasar et al., 1999; Numazaki et al., 2002), others have suggested that isoforms of PKCε (Olah et al., 2002) or PKCμ (Wang et al., 2004b) are responsible for the effects described above. Phorbol esters have also been implicated in TRPV1 activation. For example, it has been shown that a PKC-activating phorbol 12-myristate 13-acetate (PMA) decreases binding of [3H] RTX to TRPV1 (Chuang et al., 2001). Moreover, when Tyr 704 in the C-terminus is replaced with Ala, direct activation of TRPV1 by PMA is dramatically reduced (Bhave et al., 2003).

PHOSPHORYLATION BY CAMKII OF TRPV1 ACTIVITY BY CAM

AND

BINDING

AND

MODULATION

The phosphatase calcineurin inhibits desensitization of TRPV1, demonstrating that a phosphorylation/dephosphorylation process is pivotal for TRPV1 activity (Docherty et al., 1996). CaMKII controls TRPV1 activity through phosphorylation of Ser 502 and Thr 704 by regulating capsaicin binding (Jung et al., 2004). Consequently, phosphorylation of TRPV1 by three different kinases controls

TRPV1 Receptors and Signal Transduction

77

TRPV1 activity by means of an intricate balance between phosphorylation and dephosphorylation of this channel. It is known that desensitization of the TRPV1 channel depends on the presence of intracellular Ca2+, and it has recently been shown that a 35 amino acid region in the COOH-terminal region of TRPV1 (residues 767–801) binds CaM in an in vitro assay in a Ca2+-dependent manner (Numazaki and Tominaga, 2004). However, mutant channels in which this region has been deleted continued to show desensitization in whole-cell experiments, albeit with altered kinetics. In contrast, in experiments using excised membrane patches with heterologously expressed TRPV1, the application of Ca2+ with CaM produced a large reduction in current. It was also found that overexpression of CaM, together with the TRPV1 channel in this system, potentiates the inhibitory effects of Ca2+ alone, whereas coexpression of a mutant inactive form of CaM with TRPV1 does not produce a Ca2+-mediated inhibition (Rosenbaum et al., 2004). Using GST-fusion proteins corresponding to regions of the TRPV1 NH2-terminus, a binding site for CaM was localized in the region including amino acids 189–222 (Rosenbaum et al., 2004).

MODULATION

BY

LIPIDS

Membrane-derived lipids regulate the function of some ion channels, including TRPV1. For instance, TRPV1 is activated by OEA, anandamide, and some lipoxygenase products (Ahern, 2003; Hwang et al., 2000; Zygmunt et al., 1999). It has been proposed that phosphatidylinositol 4,5-bisphosphate (PIP2) is constitutively associated with TRPV1, promoting its inhibition. PIP2-mediated inhibition of TRPV1 can be released upon PLC activation by activation of metabotropic receptors and by the resulting hydrolysis of PIP2 to diacylglycerol and inositol (1,4,5) trisphosphate. Removal of PIP2 from TRPV1 by means of cleavage by PLC causes channel activation (Chuang et al., 2001). A region formed by amino acids 777–820 of TRPV1, which includes eight positively charged residues, has been proposed as a motif that regulates PIP2 (Prescott and Julius, 2003). The region in question includes Ser 800, substrates for PKC-dependent phosphorylation, and also overlaps with the C-terminus CaM-binding site. Moreover, metabolic products of LOXs, such as 12- and 15-HPETEs and 5- and 15-HETEs, are capable of activating TRPV1 (Hwang et al., 2000; see Figure 5.1B). Interestingly, bradykinin, an inﬂammatory response mediator, seems to produce nociceptor excitation by stimulating PLC, which, in turn, results in the release of inositol (1,4,5)-trisphosphate and 1,2-diacylglycerol in sensory neurons (Burgess et al., 1989; Thayer et al., 1988). Bradykinin also releases arachidonic acid (AA), a substrate for LOX, which produces excitation of sensory neurons via the phospholipase A2 (PLA2)/LOX pathway (Shin et al., 2002; Suh and Oh, 2005).

EFFECTS

OF

REDUCING AGENTS

ON

TRPV1 ACTIVITY

Agents that promote reduction or oxidation of –SH groups of cysteines inﬂuence membrane currents induced by noxious heat or capsaicin. For example, dithiothreitol (DTT), an agent that maintains –SH groups of Cys in a reduced state, facilitates

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membrane currents through TRPV1 when applied from the extracellular face of the channel by interacting with Cys located at positions 616, 621, and 634 in the loop between the ﬁfth and sixth transmembranal domains (Tousova et al., 2004). These results suggest that the sensitivity of TRPV1 to heat and capsaicin also depends on the reduced and oxidized states of the sulphydryl groups of the cysteines of the TRPV1 channel. The structural regions in TRPV1 necessary for agonist binding and modulation mentioned above are shown in Figure 5.1A. Although some regions and residues implicated in TRPV1 function have been identiﬁed, further work on the relationship between the structure and the function of this channel is required. The next few years will prove to be important in the advancement of our knowledge of the molecular mechanisms that underlie our perception of noxious stimuli.

ACKNOWLEDGMENTS This work is supported in part by DC-01065, GM-63577, GM-27278, and Philip Morris USA Inc. and Philip Morris International, and by DGAPA IN201705 and CONACyT 46004, México.

REFERENCES Acs, G., Palkovits, M., and Blumberg, P.M. (1994). Comparison of [3H]resiniferatoxin binding by the vanilloid (capsaicin) receptor in dorsal root ganglia, spinal cord, dorsal vagal complex, sciatic and vagal nerve and urinary bladder of the rat. Life Sci. 55, 1017–1026. Ahern, G.P. (2003). Activation of TRPV1 by the satiety factor oleoylethanolamide. J. Biol. Chem. 278, 30429–30434. Ahluwalia, J., Yaqoob, M., Urban, L., Bevan, S., and Nagy, I. (2003). Activation of capsaicinsensitive primary sensory neurons induces anandamide production and release. J. Neurochem. 84, 585–591. Appendino, G. and Szallasi, A. (1997). Euphorbium: modern research on its active principle, resiniferatoxin, revives an ancient medicine. Life Sci. 60, 681–696. Baumann, T.K., Burchiel, K.J., Ingram, S.L., and Martenson, M.E. (1996). Responses of adult human dorsal root ganglion neurons in culture to capsaicin and low pH. Pain 65, 31–38. Bevan, S., and Yeats, J. (1991). Protons activate a cation conductance in a subpopulation of rat dorsal root ganglion neurons. J. Physiol. (Lond) 433, 145–161. Bhave, G., Hu, H.J., Glauner, K.S., Zhu, W., Wang, H., Brasier, D.J., Oxford, G.S., and Gereau, R.W.T. (2003). Protein kinase C phosphorylation sensitizes but does not activate the capsaicin receptor transient receptor potential vanilloid 1 (TRPV1). Proc. Natl. Acad. Sci. USA 100, 12480–12485. Bhave, G., Zhu, W., Wang, H., Brasier, D.J., Oxford, G.S., and Gereau, R.W.T. (2002). cAMPdependent protein kinase regulates desensitization of the capsaicin receptor (VR1) by direct phosphorylation. Neuron 35, 721–731. Bianchi, L. and Driscoll, M. (2002). Protons at the gate: DEG/ENaC ion channels help us feel and remember. Neuron 34, 337–340.

TRPV1 Receptors and Signal Transduction

79

Brauchi, S., Orta, G., Salazar, M., Rosenmann, E., and Latorre, R. (2006). A hot-sensing cold receptor: C-terminal domain determines thermosensation in transient receptor potential channels. J. Neurosci. 26, 4835–4840. Burgess, G.M., Mullaney, I., McNeill, M., Dunn, P.M., and Rang, H.P. (1989). Second messengers involved in the mechanism of action of bradykinin in sensory neurons in culture. J. Neurosci. 9, 3314–3325. Caterina, M.J. and Julius, D. (2001). The vanilloid receptor: a molecular gateway to the pain pathway. Annu. Rev. Neurosci. 24, 487–517. Caterina, M.J., Schumacher, M.A., Tominaga, M., Rosen, T.A., Levine, J.D., and Julius, D. (1997). The capsaicin receptor: a heat-activated ion channel in the pain pathway. Nature 389, 816–824. Cesare, P., Dekker, L.V., Sardini, A., Parker, P.J., and McNaughton, P.A. (1999). Speciﬁc involvement of PKC-epsilon in sensitization of the neuronal response to painful heat. Neuron 23, 617–624. Chuang, H.H., Prescott, E.D., Kong, H., Shields, S., Jordt, S.E., Basbaum, A.I., Chao, M.V., and Julius, D. (2001). Bradykinin and nerve growth factor release the capsaicin receptor from PtdIns(4,5)P2-mediated inhibition. Nature 411, 957–962. Clapham, D.E. (2003). TRP channels as cellular sensors. Nature 426, 517–524. Cortright, D.N. and Szallasi, A. (2004). Biochemical pharmacology of the vanilloid receptor TRPV1. An update. Eur. J. Biochem. 271, 1814–1819. Costigan, M. and Woolf, C.J. (2000). Pain: Molecular mechanisms. Pain 1, 35–44. Craib, S.J., Ellington, H.C., Pertwee, R.G., and Ross, R.A. (2001). A possible role of lipoxygenase in the activation of vanilloid receptors by anandamide in the guinea-pig bronchus. Br. J. Pharmacol. 134, 30–37. Dai, Y., Moriyama, T., Higashi, T., Togashi, K., Kobayashi, K., Yamanaka, H., Tominaga, M., and Noguchi, K. (2004). Proteinase-activated receptor 2-mediated potentiation of transient receptor potential vanilloid subfamily 1 activity reveals a mechanism for proteinase-induced inﬂammatory pain. J. Neurosci. 24, 4293–4299. De Petrocellis, L., Harrison, S., Bisogno, T., Tognetto, M., Brandi, I., Smith, G.D., Creminon, C., Davis, J.B., Geppetti, P., and Di Marzo, V. (2001). The vanilloid receptor (VR1)–mediated effects of anandamide are potently enhanced by the cAMPdependent protein kinase. J. Neurochem. 77, 1660–1663. De Petrocellis, L., Orlando, P., and DiMarzo, V. (1995). Anandamide, an endogenous cannabinomimetic substance, modulates rat brain protein kinase C in vitro. Biochem. Mol. Biol. Int. 36, 1127–1133. DiMarzo, V., Blumberg, P., and Szallasi, A. (2002). Endovanilloid signalling in pain. Curr. Opinion in Neurobiol. 12, 372–379. DiMarzo, V., Fontana, A., Cadas, H., Schinellis, S., Cimino, G., Schwartz, J.C., and Pimellid, D. (1994). Formation and inactivation of endogenous cannabinoid anandamide in central neurons. Nature 372, 686–691. Docherty, R.J., Yeats, J.C., Bevan, S., and Boddeke, H.W.G.M. (1996). Inhibition of calcineurin inhibits the desensitization of capsaicin-evoked currents in cultured dorsal root ganglion neurons from adult rats. Pﬂügers Arch. 431, 828–837. Dray, A. and Dickenson, A. (1991). Systemic capsaicin and olvanil reduce the acute algogenic and the late inﬂammatory phase following formalin injection into rodent paw. Pain 47, 79–83. Farkas-Szallasi, T., Lundberg, J.M., Wiesenfeld, H.Z., Hokfelt, T., and Szallasi, A. (1995). Increased levels of GMP, VIP, and nitric oxide synthase, and their mRNAs, in lumbar dorsal root ganglia of the rat following systemic resiniferatoxin treatment. Neuroreport 6, 2230–2234.

80

TRP Ion Channel Function in Sensory Transduction

Ferrer-Montiel, A., Garcia-Martinez, C., Morenilla-Palao, C., Garcia-Sanz, N., FernandezCarvajal, A., Fernandez-Ballester, G., and Planells-Cases, R. (2004). Molecular architecture of the vanilloid receptor. Insights for drug design. Eur. J. Biochem. 271, 1820–1826. Gavva, N.R., Klionsky, L., Qu, Y., Shi, L., Tamir, R., Edenson, S., Zhang, T.J., Viswanadhan, V.N., Toth, A., Pearce, L.V., et al. (2004). Molecular determinants of vanilloid sensitivity in TRPV1. J. Biol. Chem. 279, 20283–20295. Goso, C., Evangelista, S., Tramontana, M., Manzini, S., Blumberg, P.M., and Szallasi, A. (1993). Topical capsaicin administration protects against trinitrobenzene sulfonic acid-induced colitis in the rat. Eur. J. Pharmacol. 249, 185–190. Guenther, S., Reeh, P.W., and Kress, M. (1999). Rises in [Ca2+]i mediate capsaicin- and protoninduced heat sensitization of rat primary nociceptive neurons. Europ. J. Neurosci. 11, 3143–3150. Gunthorpe, M.J., Benham, C.D., Randall, A., and Davis, J.B. (2002). The diversity in the vanilloid (TRPV) receptor family of ion channels. Trends Pharmacol. Sci. 23, 183–191. Gunthorpe, M.J., Harries, M.H., Prinjha, R.K., Davis, J.B., and Randall, A. (2000). Voltageand time-dependent properties of the recombinant rat vanilloid receptor (rVR1). J. Physiol. 525 Pt. 3, 747–759. Hellwig, N., Plant, T.D., Janson, W., Schafer, M., Schultz, G., and Schaefer, M. (2004). TRPV1 acts as proton channel to induce acidiﬁcation in nociceptive neurons. J. Biol. Chem. 279, 34553–34561. Hergenhahn, M., Kusumoto, S., and Hecker, E. (1984). On the active principles of the spurge family (Euphorbiaceae). V. Extremely skin-irritant and moderately tumor-promoting diterpene esters from Euphorbia resinifera Berg. J. Cancer Res. Clin. Oncol. 108, 98–109. Hermann, H., De Petrocellis, L., Bisogno,T., Schiano Moriello, A., Lutz, B., and DiMarzo, V. (2003). Dual effect of cannabinoid CB1 receptor stimulation on a vanilloid VR1 receptor-mediated response. Cell. Mol. Life Sci. 60, 607–616. Horie, S. Michael, G.J., and Priestley, J.V. (2005). Co-localization of TRPV1-expressing nerve ﬁbers with calcitonin-gene-related peptide and substance P in fundus of rat stomach. Inﬂammopharmacology 13, 127–137. Hui, K., Liu, B., and Qin, F. (2003). Capsaicin activation of the pain receptor, VR1: multiple open states from both partial and full binding. J. Gen. Physiol. 84, 2957–2968. Hwang, S.W., Cho, H., Kwak, J., Lee, S.Y., Kang, C.J., Jung, J., Cho, S., Min, K.H., Suh, Y.G., Kim, D., and Oh, U. (2000). Direct activation of capsaicin receptors by products of lipoxygenases: endogenous capsaicin-like substances. Proc. Natl. Acad. Sci. USA 97, 6155–6160. Jin, X., Morsy, N., Winston, J., Pasricha, P.J., Garrett, K., and Akbarali, H.I. (2004). Modulation of TRPV1 by nonreceptor tyrosine kinase, c-Src kinase. Am. J. Physiol. Cell Physiol. 287, C558–C563. Jordt, S.E. and Julius, D. (2002). Molecular basis for species-speciﬁc sensitivity to ‘‘hot’’ chili peppers. Cell 108, 421–430. Jordt, S.E., Tominaga, M., and Julius, D. (2000). Acid potentiation of the capsaicin receptor determined by a key extracellular site. Proc. Natl. Acad. Sci. USA 97, 8134–8139. Julius, D. and Basbaum, A.I. (2001). Molecular mechanisms of nociception. Nature 413, 203–210. Jung, J., Hwang, S.W., Kwak, J., Lee, S.Y., Kang, C.J., Kim, W.B., Kim, D., and Oh, U. (1999). Capsaicin binds to the intracellular domain of the capsaicin-activated ion channel. J. Neurosci. 19, 529–538.

TRPV1 Receptors and Signal Transduction

81

Jung, J., Lee, S.Y., Hwang, S.W., Cho, H., Shin, J., Kang, Y.S., Kim, S., and Oh, U. (2002). Agonist recognition sites in the cytosolic tails of vanilloid receptor 1. J. Biol. Chem. 277, 44448–44454. Jung, J., Shin, J.S., Lee, S.Y., Hwang, S.W., Koo, J., Cho, H., and Oh, U. (2004). Phosphorylation of vanilloid receptor 1 by Ca2+/calmodulin-dependent kinase II regulates its vanilloid binding. J. Biol. Chem. 279, 7048–7054. Kedei, N., Szabo, T., Lile, J.D., Treanor, J.J., Olah, Z., Iadarola, M.J., and Blumberg, P.M. (2001). Analysis of the native quaternary structure of vanilloid receptor 1. J. Biol. Chem. 276, 28613–28619. Kellenberger, S., Gautschi, I., and Schild, L. (2002). An external site controls closing of the epithelial Na+ channel ENaC. J. Physiol. 543, 413–424. Khasar, S.G., Lin, Y.H., Martin, A., Dadgar, J., McMahon, T., Wang, D., Hundle, B., Aley, K.O., Isenberg, W., McCarter, G., et al. (1999). A novel nociceptor signaling pathway revealed in protein kinase C epsilon mutant mice. Neuron 24, 253–260. Kim, S., Kang, C., Shin, C.Y., Hwang, S.W., Yang, Y.D., Shim, W.S., Park, M.Y., Kim, E., Kim, M., Kim, B.M., Cho, H., Shin, Y., and Oh, U. (2006). TRPV1 recapitulates native capsaicin receptor in sensory neurons in association with Fas-associated factor 1. J. Neurosci. 26, 2403–2412. Koplas, P.A., Rosenberg, R.L., and Oxford, G.S. (1997). The role of calcium in the desensitization of capsaicin responses in rat dorsal root ganglion neurons. J. Neurosci. 17, 3525–3537. Krause, J.E., Chenard, B.L., and Cortright, D.N. (2005). Transient receptor potential ion channels as targets for the discovery of pain therapeutics. Current Opinion in Investigational Drugs 6, 48–57. Kress, M., Fetzer, S., Reeh, P.W., and Vyklicky, L. (1996). Low pH facilitates capsaicin responses in isolated sensory neurons of the rat. Neurosci. Lett. 211, 5–8. Kuzhikandathil, E.V., Wang, H., Szabo, T., Morozova, N., Blumberg, P.M., and Oxford, G.S. (2001). Functional analysis of capsaicin receptor (vanilloid receptor subtype 1) multimerization and agonist responsiveness using a dominant negative mutation. J. Neurosci. 15, 8697–8706. Liu, L., Lo, Y., Chen, I., and Simon, S.A. (1997). The responses of rat trigeminal ganglion neurons to capsaicin and two nonpungent vanilloid receptor agonists, olvanil and glyceryl nonamide. J. Neurosci. 17, 4101–4111. Liu, L. and Simon, S.A. (1996). Similarities and differences in the currents activated by capsaicin, piperine, and zingerone in rat trigeminal ganglion cells. J. Neurophysiol. 76, 1858–1869. Liu, L. and Simon, S.A. (2000). Capsacin, acid, and heat–evoked currents in rat trigeminal ganglion neurons: relationships to functional VR1 receptors, Physiol. Behav. 69, 363–378. Lu, G., Henderson, D., Liu, L., Reinhart, P.H., and Simon, S.A. (2005). Cloning and functional characterization of TRPV1b, a new human vanilloid receptor, Molecular Pharmacology 67, 1119–1127. Maccarone, M., Salvati, S., Bari, M., and Finazzi-Agro, A. (2000). Anandamide and 2arachidonoylglycerol inhibit fatty acid amide hydrolase by activating the liproxygenase pathway of the arachidonate cascade, Biochemical and Biophysical Research Communications 278, 576–583. Mandadi, S., Numazaki, M., Tominaga, M., Bhat, M.B., Armati, P.J., and Roufogalis, B.D. (2004). Activation of protein kinase C reverses capsaicin-induced calcium-dependent desensitization of TRPV1 ion channels. Cell Calcium 35, 471–478.

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Martenson, M.E., Ingram, S.L., and Baumann, T.K. (1994). Potentiation of rabbit trigeminal responses to capsaicin in a low pH environment. Brain Res. 651, 143–147. McNamara, F.N., Randall, A., and Gunthorpe, M.J. (2005). Effects of piperine, the pungent component of black pepper, at the human vanilloid receptor (TRPV1). Br. J. Pharmacol. 144, 781–790. Mezey, E., Toth, Z.E., Cortright, D.N., Arzubi, M.K., Krause, J.E., Elde, R., Guo, A., Blumberg, P.M., and Szallasi, A. (2000). Distribution of mRNA for vanilloid receptor subtype 1 (VR1), and VR1-like immunoreactivity, in the central nervous system of the rat and human. Proc Natl Acad Sci USA 97, 3655–3660. Mohapatra, D.P. and Nau, C. (2003). Desensitization of capsaicin-activated currents in the vanilloid receptor TRPV1 is decreased by the cyclic AMP-dependent protein kinase pathway. J. Biol. Chem. 278, 50080–50090. Moriyama, T., Higashi, T., Togashi, K., Iida, T., Segi, E., Sugimoto, Y., Tominaga, T., Narumiya, S., and Tominaga, M. (2005). Sensitization of TRPV1 by EP1 and IP reveals peripheral nociceptive mechanism of prostaglandins. Mol. Pain 1, 3. Moriyama, T., Iida, T., Kobayashi, K., Higashi, T., Fukuoka, T., Tsumura, H., Leon, C., Suzuki, N., Inoue, K., Gachet, C., et al. (2003). Possible involvement of P2Y2 metabotropic receptors in ATP-induced transient receptor potential vanilloid receptor 1-mediated thermal hypersensitivity. J. Neurosci. 23, 6058–6062. Numazaki, M. and Tominaga, M. (2004). Nociception and TRP channels. Curr. Drug Targets CNS Neurol. Disord. 3, 479–485. Numazaki, M., Tominaga, T., Takeuchi, K., Murayama, N., Toyooka, H., and Tominaga, M. (2003). Structural determinant of TRPV1 desensitization interacts with calmodulin. Proc Natl Acad Sci USA 100, 8002–8006. Numazaki, M., Tominaga, T., Toyooka, H., and Tominaga, M. (2002). Direct phosphorylation of capsaicin receptor VR1 by protein kinase C epsilon and identiﬁcation of two target serine residues. J. Biol. Chem. 277, 13375–13378. Oh, U.S., Hwang, W., and Kim, D. (1996). Capsaicin activates a nonselective cation channel in cultured neonatal rat dorsal root ganglion neurons. J. Neurosci. 16, 1659–1667. Olah, Z., Karai, L., and Iadarola, M.J. (2002). Protein kinase C (alpha) is required for vanilloid receptor 1 activation. Evidence for multiple signaling pathways. J. Biol. Chem. 277, 35752–35759. Parlani, M., Conte, B., Goso, C., Szallasi, A., and Manzini, S. (1993). Capsaicin-induced relaxation in the rat isolated external urethral sphincter: characterization of the vanilloid receptor and mediation by CGRP. Br. J. Pharmacol. 110, 989–994. Patapoutian, A., Peier, A.M., Story, G.M., and Viswanath, V. (2003). ThermoTRP channels and beyond: mechanisms of temperature sensation. Nat. Rev. Neurosci. 4, 529–539. Petersen, M. and LaMotte, R.H. (1993). Effect of protons on the inward current evoked by capsaicin in isolated dorsal root ganglion cells. Pain 54, 37–42. Phillips, E., Reeve, A., Bevan, S., and McIntyre, P. (2004). Identiﬁcation of species-speciﬁc determinants of the action of the antagonist capsazepine and the agonist PPAHV on TRPV1. J. Biol. Chem. 279, 17165–17172. Piper, A.S., Yeats, J.C., Bevan, S., and Docherty, R.J. (1999). A study of the voltagedependence of capsaicin-sensitive membrane currents in rat sensory neurons before and after acute desensitization. J. Physiol. 518, 721–733. Premkumar, L.S. and Ahern, G.P. (2000). Induction of vanilloid receptor channel activity by protein kinase C. Nature 408, 985–990.

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Premkumar, L.S., Qi, Z.H., Van Buren, J., and Raisinghani, M. (2004). Enhancement of potency and efﬁcacy of NADA by PKC-mediated phosphorylation of vanilloid receptor. J. Neurophysiol. 91, 1442–1449. Prescott, E.D. and Julius, D. (2003). A modular PIP2 binding site as a determinant of capsaicin receptor sensitivity. Science 300, 1284–1288. Price, D.D. (2000). Psychological and neural mechanisms of the affective dimension of pain. Science 288, 1769–1772. Purkiss, J.R.M.J., Welch, S., Doward, K., and Foster, A. (1997). Capsaicin stimulates release of substance P from dorsal root ganglion neurons via two distinct mechanisms. Biochem. Soc. Trans. 25, 542S. Rami, H.K. and Gunthorpe, M.J. (2004). The therapeutic potential of TRPV1 (VR1) antagonists: clinical answers await. Drug Discovery Today: Therapeutic Strategies 1, 97–104. Rathee, P.K., Distler, C., Obreja, O., Neuhuber, W., Wang, G.K., Wang, S.Y., Nau, C., and Kress, M. (2002). PKA/AKAP/VR-1 module: A common link of Gs-mediated signaling to thermal hyperalgesia. J. Neurosci. 22, 4740–4745. Reeh, P.W. and Kress, M. (2002). Molecular physiology of proton transduction in nociceptors. Current Opinion in Pharmacology 1, 45–51. Rinder, J., Szallasi, A., and Lundberg, J.M. (1996). Capsaicin-, resiniferatoxin-, and lactic acid–evoked vascular effects in the pig nasal mucosa in vivo with reference to characterization of the vanilloid receptor. Pharmacol. Toxicol. 78, 327–335. Rosenbaum, T., Gordon-Shaag, A., Munari, M., and Gordon, S.E. (2004). Ca2+/calmodulin modulates TRPV1 activation by capsaicin. J. Gen. Physiol. 123, 53–62. Ross, R.A. (2003). Anandamide and vanilloid TRPV1 receptors. Br. J. Pharmacol. 140, 790–801. Ryu, S., Liu, B., and Qin, F. (2003). Low pH potentiates both capsaicin binding and channel gating of VR1 receptors. J. Gen. Physiol.122, 45–61. Schumacher, M.A., Moff, I., Samndanagunda, S.P., and Levine, J.D. (2000). Molecular cloning of an N-terminal splice variant of the capsaicin receptor. J. Biol. Chem. 275, 2756–2762. Shin, H.J., Gye, M.H., Chung, K.H., and Yoo, B.S. (2002). Activity of protein kinase C modulates the apoptosis induced by polychlorinated biphenyls in human leukemic HL-60 cells. Toxicol. Lett. 135, 25–31. Spath, E. and Darling, S.F. (1930). Synthesis of capsaicin. Ber. Chem. Ges. 63B, 737–740. Suh, Y-G. and Oh, U. (2005). Activation and activators of TRPV1 and their pharmaceutical implication. Current Pharmaceutical Design 11, 2687–2698. Szallasi, A. and Blumberg, P.M. (1999). Vanilloid (Capsaicin) receptors and mechanisms. Pharmacol. Rev. 51, 159–212. Szallasi, A. and Blumberg, P.M. (1989). Resiniferatoxin, a phorbol-related diterpene, acts as an ultrapotent analog of capsaicin, the irritant constituent in red pepper. Neuroscience 30, 515–520. Szallasi, A., Goso, C., Blumberg, P.M., and Manzini, S. (1993). Competitive inhibition by capsazepine of [3H]resiniferatoxin binding to central (spinal cord and dorsal root ganglia) and peripheral (urinary bladder and airways) vanilloid (capsaicin) receptors in the rat. J. Pharmacol. Exp. Ther. 267, 728–733. Szolcsanyi, J., Szallasi, A., Szallasi, Z., Joo, F., and Blumberg, P.M. (1990). Resiniferatoxin: an ultrapotent selective modulator of capsaicin-sensitive primary afferent neurons. J. Pharmacol. Exp. Ther. 255, 923–928. Thayer, S.A., Perney, T.M., and Miller, R.J. (1988). Regulation of calcium homeostasis in sensory neurons by bradykinin. J. Neurosci. 8, 4089–4097. Thresh, L.T. (1846). Isolation of capsaicin. Pharm J. 6, 941–947.

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Tominaga, M. and Tominaga, T. (2005). Structure and function of TRPV1. Pﬂügers Arch. 451, 143–150. Tominaga, M., Caterina, M.J., Malmberg, A.B., Rosen, T.A., Gilbert, H., Skinner, K., Raumann, B.E., Basbaum, A.I., and Julius, D. (1998). The cloned capsaicin receptor integrates multiple pain-producing stimuli. Neuron 21, 531–543. Tominaga, M., Wada, M., and Masu, M. (2001). Potentiation of capsaicin receptor activity by metabotropic ATP receptors as a possible mechanism for ATP-evoked pain and hyperalgesia. Proc. Natl. Acad. Sci. USA 98, 6951–6956. Toth, A., Boczan, J., Kedei, N., Lizanecz, E., Bagi, Z., Papp, Z., Edes, I., Csiba, L., and Blumberg, P.M. (2005). Expression and distribution of vanilloid receptor 1 (TRPV1) in the adult rat brain. Brain Res. Mol. Brain Res. 135, 162–168. Tousova, K., Susankova, K., Teisinger, J., Vyklicky, L., and Vlachova, V. (2004). Oxidizing reagent copper-o-phenanthroline is an open channel blocker of the vanilloid receptor TRPV1. Neuropharmacology 47, 273–285. Varga, A., Bolcskei, K., Szoke, E., Almasi, R., Czeh, G., Szolcsanyi, J., and Petho, G. (2006). Relative roles of protein kinase A and protein kinase C in modulation of transient receptor potential vanilloid type 1 receptor responsiveness in rat sensory neurons in vitro and peripheral nociceptors in vivo. Neuroscience 140, 645–657. Vellani, V., Mapplebeck, S., Moriondo, A., Davis, J.B., and McNaughton, P.A. (2001). Protein kinase C activation potentiates gating of the vanilloid receptor VR1 by capsaicin, protons, heat, and anandamide. J. Physiol. 534, 813–825. Vlachova, V., Teisinger, J., Susankova, K., Lyfenko, A., Ettrich, R., and Vyklicky, L. (2003). Functional role of C-terminal cytoplasmic tail of rat vanilloid receptor 1. J. Neurosci. 23, 1340–1350. Voets, T., Droogmans, G., Wissenbach, U., Janssens, A., Flockerzi, V., and Nilius, B. (2004). The principle of temperature-dependent gating in cold- and heat-sensitive TRP channels. Nature 430, 748–754. Waldmann, R., Champigny, G., Lingueglia, E., DeWeille, J.R., Heurteaux, C., and Lazdunski, M. (1999). H(+)-gated ion channels. Ann. N.Y. Acad. Sci. 868, 67–76. Walpole, C.S., Wrigglesworth, R., Bevan, S., Campbell, E.A., Dray, A., James, I.F., Masdin, K.J., Perkins, M.N., and Winter, J. (1993). Analogues of capsaicin with agonist activity as novel analgesic agents; structure-activity studies. 3. The hydrophobic sidechain ‘‘C-region.’’ J. Med. Chem. 36, 2381–2389. Wang, C., Hu, H.Z., Colton, C.K., Wood, J.D., and Zhu, M.X. (2004a). An alternative splicing product of the murine Trpv1 gene dominant negatively modulates the activity of TRPV1 channels. J. Biol. Chem. 27, 37423–37430. Wang, Y., Kedei, N., Wang, M., Wang, Q.J., Huppler, A.R., Toth, A., Tran, R., and Blumberg, P.M. (2004b). Interaction between protein kinase C mu and the vanilloid receptor type 1. J. Biol. Chem. 279, 53674–53682. Welch, J.M., Simon, S.A., and Reinhart, P.H. (2000). The activation mechanism of rat vanilloid receptor 1 by capsaicin involves the pore domain and differs from the activation by either acid or heat. Proc. Natl. Acad. Sci. USA 97, 13889–13894. Wood, J.N., Winter, J., James, I.F., Rang, H.P., Yeats, J., and Bevan, S. (1988). Capsaicininduced ion ﬂuxes in dorsal root ganglion cells in culture. J. Neurosci. 8, 3208–3220. Xue, Q., Yu, Y., Trik, S.L., Jong, B.E., and Schumacher, M.A. (2001). The genomic organization of the gene encoding the vanilloid receptor: evidence for multiple splice variants. Genomics 76, 14–20. Zygmunt, P.M., Petersson, J., Andersson, D.A., Chuang, H., Sorgard, M., Di Marzo, V., Julius, D., and Hogestatt, E.D. (1999). Vanilloid receptors on sensory nerves mediate the vasodilator action of anandamide. Nature 400, 452–457.

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Complex Regulation of TRPV1 by Vanilloids Arpad Szallasi Monmouth Medical Center and Drexel University College of Medicine

Peter M. Blumberg National Cancer Institute

CONTENTS Introduction..............................................................................................................86 The Vanilloid Receptor TRPV1 and Its Endogenous Ligands, the “Endovanilloids”................................................................................................87 TRPV1 in the Resting State ....................................................................................89 Molecular Activators of TRPV1..............................................................................89 Regulation by NGF of TRPV1 May Play a Central Role in Inﬂammatory Hyperalgesia .................................................................................90 Modulation of TRPV1 by Protein Kinases.............................................................90 Shufﬂing of TRPV1 among Various Subcellular Compartments ...........................91 Regulation of TRPV1 by Mast Cells ......................................................................91 How Can a Single Receptor Mediate Responses with Dissimilar Structure-Activity Relations? ........................................................91 Diversity of Behavior of Exogenous Ligands.........................................................92 Tissue and Cellular Expression of VR1 and Its Splice Variants ............................94 Possible Regulation of TRPV1 by Splice Variants.................................................95 Regulation of TRPV1 by Serum Steroids...............................................................95 Regulation of TRPV1 by (Neuro)endocrine Factors ..............................................95 TRPV1 May Subserve Opposing Actions at the Whole Animal Level .................96 TRPV1 Is Coexpressed with Its Relatives on Sensory Neurons with Overlapping Activity .......................................................................................96 Pharmacological Overlap between TRPV1 and Its Relatives ................................96 TRPV1 Expression May Change under Pathological Conditions..........................97 Concluding Remarks ...............................................................................................98 Acknowledgments....................................................................................................98 References................................................................................................................98

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INTRODUCTION A subset of sensory neurons is characterized by a unique sensitivity to capsaicin, the piquant ingredient in hot chili peppers.1 The excitation of these nerves by capsaicin (Figure 6.1) is followed by a lasting and fully reversible refractory state, traditionally referred to as desensitization, or, under certain conditions such as neonatal treatment, by gross neurotoxicity.1 (Parenthetically, carefully executed studies found no morphologic evidence of neurotoxicity by capsaicin at therapeutic doses.2) Capsaicin evokes these responses by interacting at a speciﬁc membrane recognition site, originally termed the vanilloid receptor.1 The diversity of capsaicin-evoked behavior at the whole animal level has, however, long puzzled scientists. For example, while a clear structure-activity relationship for capsaicin congeners was obtained in the rat eye-wiping assay, it also turned out that pungency was not proportional to the desensitizing effect.3 Based on these studies, it was postulated that different pharmacophores may be responsible for the excitatory

FIGURE 6.1 Typical naturally occurring TRPV1 agonists: capsaicin, resiniferatoxin (RTX), and evodiamine.

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and blocking actions of capsaicinoids.3 The recognition that resiniferatoxin (RTX; Figure 6.1), a diterpene ester isolated from the latex of the cactuslike plant E. resinifera, functions as an ultrapotent capsaicin analogue with a peculiar spectrum of pharmacological actions has lent further experimental support to this concept.1 For instance, in the rat RTX can desensitize the pulmonary chemoreﬂex without any apparent prior excitation, indicating that desensitization may be disconnected from stimulation.4 This is of great importance, as the initial pain response (excitation) represents the main limitation on the clinical use of vanilloids. In 1990, speciﬁc binding of [3H]RTX provided the ﬁrst direct proof for the existence of a vanilloid receptor.5 Structure-activity relationships for binding and 45Ca uptake were, however, found to be dissimilar, giving rise to the concept that these responses were mediated by functionally distinct receptors.6 The molecular cloning of the rat vanilloid receptor, subsequently renamed as the transient receptor potential vanilloid receptor 1 (TRPV1), provided the opportunity to test this hypothesis.7 It turned out that binding and 45Ca uptake were both mediated by TRPV1.8 With this discovery, the research emphasis has shifted to TRPV1 regulation as the mechanism responsible for the reality of the diversity of vanilloid actions. As discussed below, there is now mounting evidence that TRPV1 regulation is amazingly complex and is manifest at many levels, from gene expression through posttranslational modiﬁcation and formation of receptor homomers to subcellular compartmentalization and association with regulatory proteins. TRPV1 regulation is still only partially understood. Although this regulation has been reviewed exhaustively, the rapid advances in this ﬁeld necessitate frequent reevaluation of accepted concepts.

THE VANILLOID RECEPTOR TRPV1 AND ITS ENDOGENOUS LIGANDS, THE “ENDOVANILLOIDS” TRPV1 is a nonselective cation channel and is a member of the transient release potential (TRP) channel superfamily.7,9 TRPV1 helps deﬁne a subclass of these channels known as TRPV channels, which contain three conserved ankyrin domains. Several conserved protein kinase A (PKA) and protein kinase C (PKC) phosphorylation sites are also observed in TRPV1, and they have important roles in the regulation of receptor functions.7 In addition to its vanilloid sensitivity, TRPV1 is activated by noxious heat (>43°C), low pH (<6.5), a variety of inﬂammatory lipid metabolites, and phosphorylation (reviewed in references 10 and 11). Therefore, the emerging concept is that TRPV1 functions as a general sensor of noxious stimuli (a nociceptor) rather than simply as a speciﬁc membrane sensor for compounds carrying a vanillyl moiety.1,9 (As a matter of fact, this concept is nothing new because this is exactly how N. Jancsó, who almost singlehandedly transformed capsaicin from a pharmacological oddity to a widely used tool to study a speciﬁc subset of sensory neurons, had deﬁned the capsaicin receptor.12) In a much simpliﬁed way, TRPV1 can be thought of as a membrane “sensor” that is gated by heat.7,9 High noxious heat (>43°C) opens the channel on its own power, whereas some ingredients in the inﬂammatory “soup” act in concert to lower

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the heat activation threshold of TRPV1 to (below) body temperature.9,11 This latter effect is mimicked by piquant agents like capsaicin7 and piperine (black pepper),13 thus explaining why we feel these spices as “hot.” Capsaicin acts as a “gating modiﬁer” that shifts activation curves toward physiological membrane potentials.14 Interestingly, TRPV1 also causes intracellular acidiﬁcation at neutral pH via a “proton hopping permeation mechanism.”15 More generally, TRPV1 is simply a channel, and its gating is sensitive to the integrated inﬂuence of its large number of regulatory factors. Moreover, it seems clear that TRPV1’s behavior does not reﬂect the binary choice between two possible states. Rather, different ligands and regulatory factors shift its behavior in different ways. Among the newly discovered vanilloids, anandamide (N-arachidonoyl-ethanolamine)16 is no doubt the most controversial. Anandamide is an endogenous eicosanoid, which preferentially acts on cannabinoid CB1 receptors: that is, an “endocannabinoid.” Anandamide is formed “on demand” from the hydrolysis of phospholipid precursors, catalyzed by phospholipase D. Anandamide was reported to act as a full agonist at both heterologously expressed and native rat17 and human18 TRPV1. This was, however, an apparently low afﬁnity interaction, resulting in understandable reluctance in accepting anandamide as a possible “endovanilloid.” New insights, however, enhance the possibility of anandamide being a potent “endovanilloid.” Now it is highly likely that the anandamide-binding site on TRPV1 is intracellular.19 Anandamide is transported through the membrane via a speciﬁc transporter. The potency of anandamide at VR1 is enhanced by coapplication of other TRPV1 agonists such as low pH and increased temperature.20–22 Agents, like nitric oxide (NO), that stimulate the anandamide membrane transporter enhance the apparent potency of anandamide at TRPV1.23 This recognition has exciting implications. Cannabinoid CB1 and vanilloid TRPV1 receptors are coexpressed on a subset of sensory neurons.24 Generally speaking, agents that activate TRPV1 are excitatory, whereas those acting on CB1 inhibit neuronal activity.25 The binding site for anandamide on TRPV1 is intracellular;19 the anandamide recognition domain on CB1 is, in contrast, extracellular. Thus, anandamide may have opposing actions on the very same nerve terminal depending on the functional state of its membrane transporter.25 Indeed, both excitation (via TRPV1) and inhibition (via CB1) of primary sensory neurons by anandamide have been described.25 Such a mechanism may account for the bell-shaped dose-response curve for some TRPV1 agonists.26 Other “endovanilloids” were also identiﬁed. The ﬁrst example is N-arachidonoyldopamine (NADA),27 a brain substance that is similar to capsaicin not only structurally but also with regard to its potency at TRPV1. Certain lipoxygenase metabolites, such as 12-HPETE, have also been identiﬁed as putative endogenous TRPV1 agonists.28 These ﬁndings may provide a mechanistic link between TRPV1 and the 5-lipoxygenase products that are important mediators of airway inﬂammation. Ethanol has also been added to the list of controversial vanilloids.29 High concentrations of ethanol activate C-ﬁbers innervating the esophagus and the skin, evoking neuropeptide release and resultant neurogenic inﬂammation.29 Ethanol also excites dorsal root ganglion neurons possessing native vanilloid receptors as well

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FIGURE 6.2 TRPV1 activation represents the balance between factors favoring its open (active) and closed (resting) states.

as HEK cells transfected with TRPV1.29 This excitation by ethanol is abolished by the coadministration of capsazepine (Figure 6.2). Even more interesting, ethanol lowers the heat threshold of TRPV1 activation from 42°C to 34°C.29

TRPV1 IN THE RESTING STATE There is mounting evidence that TRPV1 is balanced on the edge between open and closed states (Figure 6.2). Agents that promote the open state are nociceptive.11,25 By contrast, agents that shift TRPV1 toward the closed state are antinociceptive.11 As of today, phosphatidylinositol(4,5)-bisphosphate (PIP2) is the only known endogenous ligand that helps keep TRPV1 in the closed state.30 Functional recovery from desensitization of TRPV1 depends on the replenishment of PIP2.31 The inhibitory control of PIP2 is facilitated by adenosine.32 By contrast, ATP activates TRPV1.33 Of course, to the degree that TRPV1 is sensitized by phosphorylation by its many regulatory kinases such as PKC or PKA, the activity of protein phosphatases such as calcineurin will exert an inhibitory, antinociceptive effect.34

MOLECULAR ACTIVATORS OF TRPV1 Agents that favor the open state of TRPV1 are numerous and can be divided into two major categories: (1) agents that increase the probability of channel opening directly, and (2) agents that release TRPV1 from the inhibitory control of PIP2.

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The ﬁrst category is exempliﬁed by the enzymes PKC and PKA (reviewed in reference 11). Factors that stimulate phospholipase C, the enzyme that cleaves PIP2, fall into the second category.30 However, this category is typically not independent of the ﬁrst, because the breakdown of PIP2 by phospholipase C is linked to formation of diacylglycerol, the endogenous activator for PKC.35 Bradykinin (acting on B2 receptors) thus has a dual effect on TRPV1: it stimulates PKC by promoting diacylglycerol formation and, at the same time, it degrades PIP2 by facilitating phospholipase C.30 It is tempting to speculate that this dual action on TRPV1 gives an important contribution to the profound algesic activity of bradykinin. Nerve growth factor (NGF), a key player in inﬂammatory hyperalgesia, also releases TRPV1 under the inhibitory control of PIP2.30

REGULATION BY NGF OF TRPV1 MAY PLAY A CENTRAL ROLE IN INFLAMMATORY HYPERALGESIA In addition to sensitizing TRPV1 by cleaving PIP2,30 NGF may also regulate the transcription of the TRPV1 gene. It has long been known that neuropeptides are depleted from sensory neurons following capsaicin desensitization, and it was speculated that this effect is secondary to the disruption by capsaicin of centripetal axonal NGF transport from the periphery to the perikarya (reviewed in reference 1). Because the loss and recovery of speciﬁc RTX binding sites parallel the changes in neuropeptide expression,36 it is not unlikely that NGF is also required for the transcription of the TRPV1 gene. The action of NGF on TRPV1 expression may even be bidirectional. It is well documented that NGF levels are elevated under inﬂammatory conditions37 where TRPV1 is overexpressed both in animal models and human disease states. Studies with TRPV1-deﬁcient (−/−) mice have furnished experimental proof that TRPV1 is instrumental in the development and maintenance of certain types of inﬂammatory hyperalgesia.38,39 Thus, it may be speculated that NGF may play a pivotal role in causing inﬂammatory hyperalgesia by elevating TRPV1 levels40 and sensitizing the receptor30 at the same time. If so, the pharmacological manipulation of NGF–TRPV1 interactions may represent a novel therapeutic approach for the relief of inﬂammatory pain.

MODULATION OF TRPV1 BY PROTEIN KINASES PKC is central to TRPV1 regulation inasmuch as it couples an array of receptors to TRPV1 (reviewed in reference 11). Some examples were discussed above. Other notable examples include the chemokine receptor CCR1,41 the metabotropic purinergic receptor P2Y,42 and the prostaglandin receptors EP1 and IP.43 PKA and PKC are, however, not the only kinases to regulate TRPV1. The Ca2+/calmodulin-dependent kinase II (CaMKII) sensitizes TRPV1 by phosphorylation44 as does phophatidylinositol 3-kinase (PI3K) via its downstream target AKT.45 This latter ﬁnding links TRPV1 to the ERK (extracellular signal-regulated protein

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kinase) pathway. The nonreceptor tyrosine kinase Src likewise potentiates capsaicin-induced currents.46

SHUFFLING OF TRPV1 AMONG VARIOUS SUBCELLULAR COMPARTMENTS TRPV1 appears to be mostly sequestered in intracellular compartments where it exists in a homomeric complex, most likely as a tetramer.10 In fact, a tetramerization domain is present in the C-terminus of TRPV1.47 Formation of heteromers with related TRP channels like TRPV2 was also described.48 Moreover, TRPV1 is believed to be associated with cytoplasmic proteins that can further ﬁne-tune the channel’s activity. An example of this phenomenon is interaction with β-tubulin, for which TRPV1 carries a binding domain on its C-terminus.49 Upon activation, neurons begin trafﬁcking TRPV1 to the membrane, where this receptor gets activated, desensitized, and then “recycled” to the intracellular compartments. Translocation of TRPV1 to the cell membrane occurs via SNARE (snapin and synaptotagmin IX)-mediated exocytosis.50 Broadly speaking, activation involves phosphorylation by protein kinases (both PKA and PKC), and desensitization involves dephosphorylation by phosphatases (e.g., calcineurin; reviewed in references 10 and 11).

REGULATION OF TRPV1 BY MAST CELLS An added level of TRPV1 regulation is by inﬂammatory cells such as mast cells. Mast cells release tryptase that, in turn, activates the protease-activated receptor PAR-2; activation of PAR-2 then opens TRPV1 via PKC.51 In keeping with this, PAR-2 agonists reduce the heat activation threshold of TRPV1 from 42°C to below body temperature.52 Excited nerve endings release SP that, as a positive feedback, binds to neurokinin NK1 receptors on mast cells. Mast cells also express TRPV1.53 A relevant ﬁnding is that PAR-2 is upregulated in the bladder during experimental cystitis.54

HOW CAN A SINGLE RECEPTOR MEDIATE RESPONSES WITH DISSIMILAR STRUCTUREACTIVITY RELATIONS? A combination of pharmacokinetics and differential receptor regulation may explain the strikingly different structure-activity relations for vanilloid-induced Ca2+-uptake versus inhibition of [3H]RTX binding. Recently, no fewer than ﬁve parameters (potency, maximal response, latency of response, variability in latency, and desensitization) were identiﬁed in which TRPV1 agonists differ even in a simple in vitro assay like calcium response in CHO cells transfected with rTRPV1.55 For the most part, the correlation between such in vitro parameters and in vivo responses has not been deﬁned.

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The mapping of pharmacologically “hot” points that may account for this differential regulation is in progress. For example, Thr370 is the key residue that is phosphorylated by PKC (sensitization) and dephosphorylated by calcineurin (desensitization).56 Other residues phosphorylated by PKC include Ser502 and Ser800.57 High afﬁnity [3H]RTX binding has been linked to a single residue, Met547, in the S4 membrane domain, which is believed to form a “binding pocket” with Tyr511 in the S3 domain.58 It is also possible that binding and Ca2+ uptake detect two distinct populations of TRPV1: binding may predominantly detect receptors sequestered in intracellular compartments, whereas the Ca2+-uptake response reﬂects activated TRPV1, transported into the plasma membrane. As these receptor populations are in different subcellular milieu, their pharmacology can be distinct.

DIVERSITY OF BEHAVIOR OF EXOGENOUS LIGANDS The rapid advances in the characterization of TRPV1 have been matched by intense efforts to develop therapeutics targeted to TRPV1. These medicinal chemistry efforts have again revealed great diversity. From an early concept that the vanilloid moiety shared by capsaicin and RTX (compare structures in Figure 6.1) was a critical feature of ligands for the “vanilloid receptor,”1 it is now apparent that a wide variety of structures can interact with high afﬁnity. Likewise, the initial identiﬁcation of vanilloid agonists such as capsaicin, RTX, and evodiamine is now complemented by an impressive variety of potent antagonists for TRPV1, acting competitively with capsaicin (reviewed in reference 59; for selected structures, see Figure 6.3). Such antagonists functionally display at least two different behavior patterns. Some are competitive with capsaicin but are less effective for antagonizing TRPV1 activation by other regulators such as pH and temperature. Others block all three responses. A rationale for this difference is that the former may simply occupy the capsaicinbinding site, whereas for the latter class occupancy is coupled to stabilization of the closed conformation of the channel (Figure 6.4). Between antagonists and agonists are positioned another class of ligands: competitive partial agonists/partial antagonists (Figure 6.4). Although this class has received relatively little attention, such compounds may be of particular potential interest because the ratio of agonism to antagonism is not ﬁxed but rather is subject to modulation by the regulatory environment in which TRPV1 is found (Figure 6.5). It may be possible to design ligands that only function as agonists and thereby induce desensitization only in a speciﬁc environment. If so, they could represent agents that might be administered systemically to achieve a localized effect. Such compounds illustrate the potential of diversity within the TRPV1 environment to control efﬁcacy of ligands (Figure 6.5). The evidence for different structureactivity relations for different vanilloid responses also implies strongly that the TRPV1 environment can play a major role in determining structure-activity relations for potency. Although this concept does not appear to have been factored into current screening of the output from programs in medicinal chemistry, it again suggests that it may be possible to optimize vanilloid design not simply for TRPV1 but rather for TRPV1 as immersed in a speciﬁc regulatory environment (Figure 6.5).

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FIGURE 6.3 Selected examples of TRPV1 antagonists: capsazepine, iodo-RTX, BCTC, and N-arylcinnamides (AMG 9810 analogue).

FIGURE 6.4 The diverse mechanisms of action of TRPV1 ligands.

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FIGURE 6.5 Opportunities for modulating TRPV1 by interfering with its environmental regulation.

TISSUE AND CELLULAR EXPRESSION OF VR1 AND ITS SPLICE VARIANTS TRPV1 is expressed in small, unmyelinated sensory nerve ﬁbers called C ﬁbers, which conduct very slow action potentials and contain various neuropeptides including CGRP and SP (reviewed in reference 1). These TRPV1 immunoreactive positive ﬁbers have been observed innervating the skin, the bladder, and the gastric mucosa, just to cite a few examples. TRPV1 RNA and immunoreactivity are also found in the central nervous system, albeit at much lower levels than found in dorsal root ganglia (reviewed in reference 60). There is good evidence for TRPV1 gene expression in other nonneural tissues as well, including human skin (keratinocytes, dermal blood vessels, hair follicles, sebocytes, and sweat glands53,61), the bladder (urothelium, smooth muscle, mast cells, and capillaries62), and the brain microvasculature endothelium.63 TRPV1 in these tissues is functional, lending experimental foundation to the novel concept that both squamous and transitional epithelia can play sensory roles.64 In animals, the tissue distribution of TRPV1 is even broader, encompassing hearts, livers, kidneys, spleens, and lungs.65–68 Of note, rat cardiomyocytes express TRPV1 in neonates, but not in adults, suggesting that TRPV1 expression is developmentally regulated, at least in the heart.65 The identiﬁcation of CNS and peripheral nonneuronal sites of TRPV1 expression negates one of the central dogmas of the ﬁeld (that is, vanilloid receptor expression is a functional signature of primary sensory neurons) and should revolutionize the ways we think about vanilloid receptors.

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POSSIBLE REGULATION OF TRPV1 BY SPLICE VARIANTS Several splice variants of VR1 have been reported. One that is called the rat TRPV1 5’ splice variant (VR1 5’sv) differs from TRPV1 only in the amino terminal tail region.69 A similar, but not identical, variant has also been isolated from mice and is called TRPV1b.70 Neither VR1 5’sv nor TRPV1b is sensitive to capsaicin or low pH; interestingly, TRPV1b may be activated by high temperature.70 Given the hypothesis that TRPV1 is a tetramer, it is tempting to speculate that these splice variants may form heteromeric channels with wild-type TRPV1, and they may function as dominant negative regulators. VR1 5’sv RNA is expressed at very low levels in sensory neurons relative to TRPV1 but is comparable to TRPV1 gene expression in the CNS.66

REGULATION OF TRPV1 BY SERUM STEROIDS Several endogenous steroids can also bind and inhibit TRPV1. For example, dehydroepiandrosterone (DHEA), a major blood steroid, can reversibly inhibit capsaicininduced currents in dorsal root ganglion neurons with an EC50 value of 6.7 μM,71 which is close to the physiological concentration of this compound. Because DHEA levels climax in the mid-twenties and then decrease with age, elderly people might be more sensitive to capsaicin than young adults. Even more interesting, distinct structure-activity relationships were discovered for steroids: for instance, 3epiDHEA could potentiate, and not inhibit, capsaicin responses,71 raising the possibility that the steroid framework might provide an interesting platform for the discovery of new TRPV1 antagonists. At a molecular level, it is not clear if steroids are allosteric modulators of TRPV1 or if they bind directly to the capsaicin-binding domain of the receptor. For example, pregnenolone appears to have a noncompetitive mechanism.72 Another intriguing observation is that the female sex hormone 17-estradiol could dramatically potentiate capsaicin responses, whereas the male hormone testosterone had marginal inhibitory activity.73 Sex differences in pain responses have long been known, with women being more sensitive to capsaicin-induced pain than men.74 The differential modulation of capsaicin responses by female and male hormones might explain this observation.

REGULATION OF TRPV1 BY (NEURO)ENDOCRINE FACTORS There is a growing body of evidence that TRPV1 is also subject to (neuro)endocrine regulation. An interesting example of this phenomenon is insulin. TRPV1 is expressed on pancreatic islet cells where it is involved in insulin release.75 Circulating insulin, in turn, may promote translocation of TRPV1 from cytosol to plasma membrane in neurons, leading to sensitization.

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TRPV1 MAY SUBSERVE OPPOSING ACTIONS AT THE WHOLE ANIMAL LEVEL Recently, chronic mechanical hyperalgesia was compared in wild-type (WT) and TRPV1-deﬁcient mice in animal models of human diabetic (streptozoticin-induced) and toxic polyneuropathic (cisplatin-evoked) pain. Surprisingly, TRPV1 knockout mice fared worse: hyperalgesia both developed earlier and was more severe in TRPV1-deﬁcient mice compared to WT animals.76 By contrast, no difference was found between the two groups following carrageenan treatment or mechanical injury.76 Two alternative, but not mutually exclusive, mechanisms can account for the protective role of TRPV1 against certain sources of hyperalgesia. First, activation of TRPV1 by endovanilloids may release endogenous analgesic substances like somatostatin.76 Second, such endovanilloids that possess analgesic actions can be generated.77

TRPV1 IS COEXPRESSED WITH ITS RELATIVES ON SENSORY NEURONS WITH OVERLAPPING ACTIVITY TRPV1 has a number of close relatives (belonging to the extended family of TRP receptors): some (similarly to TRPV1) are involved in sensory reception, whereas others have unknown functions. Curiously, the “cool” menthol receptor transient receptor potential melastatin subfamily member 8 (TRPM8)78 and the noxious cold receptor transient receptor potential subfamily A (ankyrin-like) member 1 (TRPA1)79 are close relatives of the “hot” capsaicin receptor TRPV1. It is known that mustard oil, traditionally used as a pungent activator of capsaicin-sensitive nerves, in fact acts through TRPA1,79 providing a rationale to explain how noxious cold can paradoxically be perceived as burning pain. Of note, both TRPV130 and TRPA179 are downstream targets for bradykinin.

PHARMACOLOGICAL OVERLAP BETWEEN TRPV1 AND ITS RELATIVES TRPM8 shares many functional and pharmacological properties with TRPV1: for instance, both receptors are under the inhibitory control of PIP2.30,78 The pharmacological overlap between TRPM8 and TRPV1 is extensive80 and includes the “selective TRPV1 antagonists” N-(4-tertiarybutylphenyl)-4-(3-chloropyridin-2-yl) tetrahydropyrazine-1(2H)-carboxamide (BCTC; Figure 6.3), (2R)-4-(3-chloro-2pyridinyl)-2-methyl-N-(4-[triﬂuoromethyl]phenyl)-1-piperazinecarboxamide (CTPC), and N-(2-bromophenyl)-N’-(2-[ethyl{3-methylphenyl}amino]ethyl)-urea (SB-452533). TRPM8 and TRPV1 are colocalized in many tissues, including prostate cancer cells.81 It was postulated that TRPM8 is required for cell survival,82 whereas TRPV1 promotes apoptosis.83 If this hypothesis holds true, it is impossible

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to predict whether an antagonist that blocks both TRPM8 and TRPV1 would inhibit or accelerate prostate cancer growth.

TRPV1 EXPRESSION MAY CHANGE UNDER PATHOLOGICAL CONDITIONS Several lines of evidence demonstrate that TRPV1 protein levels are regulated. For instance, increased TRPV1 immunoreactive ﬁber innervation has been observed in inﬂamed human skin,84 vulvas,85 and gastrointestinal tracts (both in the esophagus86 and large intestine87). This phenomenon is believed to contribute to the pathogenesis of various diseases, such as reﬂux esophagitis (also known as GERD, or gastroesophageal reﬂux disease), inﬂammatory bowel disease (both Crohn’s disease and ulcerative colitis), irritable bowel syndrome, vulvar allodynia, and prurigo nodularis (reviewed in reference 88). At the cellular level, both up- and downregulation of the TRPV1 gene have been demonstrated. In the rat, reduced levels of TRPV1 mRNA were found following vanilloid desensitization89 in keeping with the loss of receptor protein (speciﬁc [3H]RTX binding) in these animals.36 By contrast, elevated levels of TRPV1 mRNA and protein were shown in animal models of inﬂammatory hyperalgesia.90 The latter ﬁnding is in agreement with the increase in TRPV1-like immunoreactivity detected in painful human disease conditions.84–88 The molecular mediators that regulate the TRPV1 gene are yet to be explored but, as discussed above, NGF is an important candidate molecule. A new ﬁnding with far-reaching implications is the demonstration of “aberrant” TRPV1 expression in cervical carcinoma cells,91 implying a future role for TRPV1 agonists as adjuvant chemotherapeutic drugs in cancer treatment regimens. The presence of neuronal capsaicin receptors in human airways is long established based on both functional studies and RTX binding experiments, but it is a recent recognition that lung epithelial cells and bronchial smooth muscle also express TRPV1.92 Capsaicin evokes the cough reﬂex in human volunteers, and the cough response is exaggerated in patients with asthma or chronic obstructive pulmonary disease.93 Recently, increased levels of TRPV1 were found in airways smooth muscle of patients with chronic cough.94 Interestingly, the increased TRPV1 immunoreactivity in skin nerve ﬁbers of women with breast pain is accompanied by elevated TRPV3 (the camphor receptor) and TRPV4 levels in keratinocytes.84 TRPV3-deﬁcient mice show a similar phenotype to TRPV1 knockout mice, including lack of inﬂammatory hyperalgesia to heat.95 Postulating that epidermal TRPV3 and TRPV4 contribute to mastalgia, an antagonist that blocks all these three receptors may have increased efﬁcacy in relieving breast pain. An experimental support for this hypothesis is provided by anandamide, an analgesic endocannabinoid, that activates TRPV1 directly and TRPV4 indirectly via a metabolite.

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CONCLUDING REMARKS Vanilloid TRPV1 receptor agonists like capsaicin and RTX are drugs with proven therapeutic value; their clinical utility is limited by a combination of the initial pain response that they evoke and their poor bioavailability. The current interest in developing TRPV1 ligands into clinically useful drugs is heavily biased toward antagonists. Whereas, at least in principle, TRPV1 antagonists should be devoid of the undesired side effects of agonists, the recent recognition of functional TRPV1 in brain nuclei as well as in a broad array of nonneuronal tissues predicts that longterm administration of TRPV1 antagonists might also lead to complications. For instance, it was postulated that TRPV1-expressing nerves are instrumental in the maintenance of normal blood pressure.96 If this hypothesis holds true, prolonged use of TRPV1 antagonists may lead to hypertension or may aggravate preexistent disease. The biochemical regulation of TRPV1 is, however, complex (Figures 6.2 and 6.5). This discovery implies that TRPV1 should be studied in its speciﬁc regulatory environment that may be disease state–dependent. The ultimate goal would be the synthesis of ligands that speciﬁcally target TRPV1 in diseased (e.g., inﬂamed) tissues but spare TRPV1 that subserves its physiological functions in healthy tissues.

ACKNOWLEDGMENTS This research was supported in part by the NIH, National Cancer Institute, Center for Cancer Research.

REFERENCES 1. Szallasi, A. and Blumberg, P.M. Vanilloid (capsaicin) receptors and mechanisms. Pharmacol. Rev. 51: 159, 1999. 2. Avelino, A. and Cruz, F. Peptide immunoreactivity and ultrastructure of rat urinary bladder nerve ﬁbers after topical desensitization by capsaicin or resiniferatoxin. Auton. Neurosci. 86: 37, 2000. 3. Szolcsányi, J. Forty years in capsaicin research for sensory pharmacology and physiology. Neuropeptides 38: 377, 2004. 4. Szolcsányi, J., Szallasi, A., Szallasi, Z., Joó, F., and Blumberg, P.M. Resiniferatoxin, an ultrapotent selective modulator of capsaicin-sensitive primary afferent neurons. J. Pharmacol. Exp. Ther. 255: 923, 1990. 5. Szallasi, A. and Blumberg, P.M. Speciﬁc binding of resiniferatoxin, an ultrapotent capsaicin analog, by dorsal root ganglion membranes. Brain Res. 524: 106, 1990. 6. Ács, G., Lee, J., Marquez, V., and Blumberg, P.M. Distinct structure-activity relations for stimulation of 45Ca uptake and for high afﬁnity binding in cultured rat dorsal root ganglion neurons and dorsal root ganglion membranes. Brain Res. Mol. Brain Res. 35: 173, 1996. 7. Caterina, M.J., Schumacher, M.A., Tominaga, M., Rosen, T.A., Levine, J.D., and Julius, D. The capsaicin receptor: a heat-activated ion channel in the pain pathway. Nature 389: 816, 1997.

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8. Szallasi, A., Blumberg, P.M., Annicelli, L.L., Krause, J.E., and Cortright, D.N. The cloned vanilloid receptor VR1 mediates both R-type binding and C-type calcium response in dorsal root ganglion neurons. Mol. Pharmacol. 56: 581, 1999. 9. Caterina, M. and Julius, D. The vanilloid receptor: a molecular gateway to the pain pathway. Annu. Rev. Neurosci. 24: 487, 2001. 10. Cortright, D.N. and Szallasi, A. Biochemical pharmacology of the vanilloid receptor TRPV1. An update. Eur. J. Biochem. 271: 1814, 2004. 11. Di Marzo, V., Blumberg, P.M., and Szallasi, A. Endovanilloid signaling in pain. Curr. Opin. Neurobiol. 12: 372, 2000. 12. Mr. and Mrs. Jancsó, M. Desensitization of sensory nerve endings (in Hungarian). Kísèrletes Orvostudomány (Experimental Medicine), 2: 15, 1949 [English translation in Ref. 1] 13. McNamara, F.N., Randall, A., and Gunthorpe, M.J. Effects of piperine, the pungent component of black pepper, at the human vanilloid receptor VR1 (TRPV1). Br. J. Pharmacol. 144: 781, 2005. 14. Voets, T., Droogmans, G., Wissenbach, U., Janssens, A., Flockerzi, V., and Nilius, B. The principle of temperature-dependent gating in cold- and heat-sensitive TRP channels. Nature 430: 748, 2002. 15. Hellwig, N., Plant, T.D., Janson, W., Schafer, M., Schultz, G., and Schaefer, M. TRPV1 acts as proton channel to induce acidiﬁcation in nociceptive neurons. J. Biol. Chem. 279: 34553, 2004. 16. Zygmunt, P.M., Petersson, J., Andersson, D.A., Chuang, H-H., Sørgård, M., DiMarzo, V., Julius, D., and Högestatt, T. Vanilloid receptors on sensory nerves mediate the vasodilator action of anandamide. Nature 400: 452, 1999. 17. Tognetto, M., Amadesi, S., Harrison, S., Creminon, C., Trevisani, M., Carreras, M., Matera, M., Geppetti, P., and Bianchi, A. Anandamide excites central terminals of dorsal root ganglion neurons via vanilloid receptor-1. J. Neurosci. 21: 1104, 2001. 18. Smart, D., Gunthorpe, M.J., Jerman, J.C., Nasir, S., Gray, J., Muir, A.I., Chambers, J.K., Randall, A.D., and Davis, J.B. The endogenous lipid anandamide is a full agonist at the human vanilloid receptor (hVR1). Br. J. Pharmacol. 129: 227, 2000. 19. Gavva, N.R., Klionsky, L., Qu, Y., Shi, L., Tamir, R., Edenson, S., Zhang, T.J., Viswanadhan, V.N., Toth, A., Pearce, L.V., Vanderah, T.W., Porreca, F., Blumberg, P.M., Lile, J., Sun, Y., Wild, K., Louis, J-C., and Treanor, J.J.S. Molecular determinants of vanilloid sensitivity in TRPV1. J. Biol. Chem. 279: 20283, 2004. 20. Premkumar, L.S. and Ahern, G.P. Induction of vanilloid receptor channel activity by protein kinase C. Nature 408: 985, 2000. 21. Vellani, V., Mapplebeck, S., Moriondo, A., Davis, J.B., and McNaughton, P.A. Protein kinase C activation potentiates gating of the vanilloid receptor VR1 by capsaicin, protons, heat, and anandamide. J. Physiol. 534: 813, 2001. 22. Olah, Z., Karai, I., and Iadarola, M. Anandamide activates vanilloid receptor (VR1) at acidic pH in dorsal root ganglia and cells ectopically expressing VR1. J. Biol. Chem. 276: 31163, 2001. 23. De Petrocellis, L., Bisogno, T., Macarrone, M., Davis, J.B., Finazzi-Agro, A., and DiMarzo, V. The activity of anandamide at vanilloid VR1 receptors requires facilitated transport across the cell membrane and is limited by intracellular metabolism. J. Biol. Chem. 276: 12856, 2001. 24. Ahluwalia, J., Urban, L., Capogna, M., Bevan, S., and Nagy, I. Cannabinoid 1 receptors are expressed on nociceptive primary sensory neurons. Neuroscience 100: 685, 2000.

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25. DiMarzo, V., Bisogno, T., and De Petrocellis, L. Anandamide: some like it hot. Trends Pharmacol. Sci. 22: 346, 2001. 26. Nèmeth, H., Helyes, Z., Than, M., Jakab, B., Pintèr, E., and Szolcsányi, J. Concentration-dependent dual effect of anandamide on sensory neuropeptide release from isolated rat tracheae. Neurosci. Lett. 336: 89, 2003. 27. Huang, S.M., Bisogno, T., Trevisani, M., Al-Hayani, A., De Petrocellis, L., Fezza, F., Tognetto, M., Petros, T.J., Krey, J.F., Chu, C.I., Miller, J.D., Davies, S.N., Geppetti, P., Walker, J.M., and DiMarzo, V. An endogenous capsaicin-like substance with high potency at recombinant and native vanilloid VR1 receptors. Proc. Natl. Acad. Sci. USA 99: 8400, 2002. 28. Hwang, S.W., Cho, J., Kwak, L., Lee, S.Y., Kang, J., Jung, S., Cho, K.H., Min, Y.G., Suh, D., Kim, U., and Oh, U. Direct activation of capsaicin receptors by products of lipoxygenases: endogenous capsaicin-like substances. Proc. Natl. Acad. Sci. USA 97: 6155, 2000. 29. Trevisani, M., Smart, D., Gunthorpe, M.J., Tognetto, M., Barbieri, M., Campi, B., Amadesi, S., Gray, J., Jerman, J.C., Brough, S.J., Owen, D., Smith, G.D., Randall, A.D., Harrison, S., Bianchi, A., Davis, J.B., and Geppetti, P. Ethanol elicits and potentiates nociceptor responses via the vanilloid receptor-1. Nat. Neurosci. 5: 546, 2002. 30. Chuang, H.H., Prescott, E.D., Kong, H., Shields, S., Jordt, S.E., Basbaum, A., Chao, M.V., and Julius, D. Bradykinin and nerve growth factor release the capsaicin receptor from PtdIns(4,5)P2-mediated inhibition. Nature 411: 957, 2001. 31. Liu, B., Zhang, C., and Qin, F. Functional recovery from desensitization of vanilloid receptor TRPV1 requires resynthesis of phosphatidylinositol 4,5-bisphosphate. J. Neurosci. 25: 4835, 2005. 32. Puntambekar, P., Van Buren, J., Raisinghani, M., Premkumar, L.S., and Ramkumar, V. Direct interaction of adenosine with the TRPV1 channel protein. J. Neurosci. 24: 3663, 2004. 33. Lakshmi, S. and Joshi, P.G. Co-activation of P2Y2 receptor and TRPV channel by ATP: implications for ATP-induced pain. Cell. Mol. Neurobiol. 25: 819, 2005. 34. Jung, J., Shin, J.S., Lee, S.Y., Hwang, S.W., Koo, J., Cho, H., and Oh, U. Phosphorylation of vanilloid receptor 1 by Ca2+/calmodulin-dependent kinase II regulates its vanilloid binding. J. Biol. Chem. 279: 7048, 2004 35. Yang, C. and Kazanietz, M.G. Divergence and complexities in DAG signaling: looking beyond PKC. Trends Pharmacol. Sci. 24: 602, 2003. 36. Szallasi, A. and Blumberg, P.M. Vanilloid receptor loss in rat sensory ganglia associated with long-term desensitization to resiniferatoxin. Neurosci. Lett. 136: 51, 1992. 37. Shu, X.Q. and Mendell, L.M. Neurotrophins and hyperalgesia. Proc. Natl. Acad. Sci. USA 96: 7693, 1999. 38. Davis, J.B., Gray, J., Gunthorpe, M.J., Hatcher, J.P., Davey, P.T., Harries, P., Harries, M.H., Latcham, J., Clapham, C., Atkinson, K., Rance, S.A., Grau, E., Harper, A.J., Pugh, P.L., Rogers, D.C., Randall, S., Randall, A., and Sheardown, S.A. Vanilloid receptor-1 is essential for inﬂammatory thermal hyperalgesia. Nature 405: 183, 2000. 39. Caterina, M.J., Lefﬂer, A., Malmberg, A.N., Martin, W.J., Trafton, J., Petersen-Zeitz, K.R., Koltzenburg, M., Basbaum, A.I., and Julius, D. Impaired nociception and pain sensation in mice lacking the capsaicin receptor. Science 288: 306, 2000. 40. Amaya, F., Shimosato, G., Nagano, M., Ueda, M., Hashimoto, S., Tanaka, Y., Suzuki, M., and Tanaka, M. NGF and GDNF differentially regulate TRPV1 expression that contributes to development of inﬂammatory hyperalgesia. Eur. J. Neurosci. 20: 2303, 2004.

Complex Regulation of TRPV1 by Vanilloids

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41. Zhang, N., Inan, S., Cowan, A., Sun, R., Wang, J.M., Rogers, T.J., Caterina, M., and Oppenheim, J.J. A proinﬂammatory chemokine, CCL3, sensitizes the heat- and capsaicin-gated ion channel TRPV1. Proc. Natl. Acad. Sci. USA 102: 4536, 2005. 42. Tominaga, M., Numazaki, M., Iida, T., Moriyama, T., Togashi, K., Higashi, T., Murayama, N., and Tominaga, T. Regulation mechanisms of vanilloid receptors. Novartis Found. Symp. 261: 4, 2004. 43. Moriyama, T., Higashi, T., Togashi, K., Iida, T., Segi, E., Sugimoto, Y., Tominaga, T., Naruyima, S., and Tominaga, M. Sensitization of TRPV1 by EP1 and IP reveals peripheral nociceptive mechanisms of prostaglandins. Mol. Pain 1: 3, 2005. 44. Jung, J., Shin, J.S., Lee, S.Y., Hwang, S.W., Koo, J., Cho, H., and Oh, U. Phosphorylation of vanilloid receptor 1 by Ca2+/calmodulin-dependent kinase II regulates its vanilloid binding. J. Biol. Chem. 279: 7048, 2004. 45. Zhuang, Z.Y., Xu, H., Clapham, D.E., and Ji, R.R. Phosphatidylinositol 3-kinase activates ERK in primary sensory neurons and mediates inﬂammatory hyperalgesia through TRPV1 sensitization. J. Neurosci. 24: 8300, 2004. 46. Jin, X., Morsy, N., Winston, J., Pasricha, P.J., Garrett, K., and Akbarali, H.I. Modulation of TRPV1 by nonreceptor kinase, c-Src kinase. Am. J. Physiol. Cell Physiol. 287: 558, 2004. 47. Garcia-Sanz, N., Fernandez-Carjaval, A., Morenilla-Palao, C., Planells-Cases, R., Fajardo-Sanchez, E., Fernandez-Ballaster, G., and Ferrer-Montiel, A. Identiﬁcation of a tetramerization domain in the C-terminus of the vanilloid receptor. J. Neurosci. 24: 5307, 2004. 48. Rutter, A.R., Ma, Q.P., Leveridge, M., and Bonnert, T.P. Heteromerization and colocalization of TrpV1 and TrpV2 in mammalian cell lines and rat dorsal root ganglia. NeuroReport 16: 1735, 2005. 49. Goswami, C., Dreger, M., Jahnel, R., Bogen, O., Gillen, C., and Hucho, F. Identiﬁcation and characterization of a Ca2+-sensitive interaction of the vanilloid receptor TRPV1 with tubulin. J. Neurochem. 91: 1092, 2004. 50. Morenilla-Palao, C., Planells-Cases, R., Garcia-Sanz, N., and Ferrer-Montiel, A. Regulated exocytosis contributes to protein kinase C potentiation of vanilloid receptor activity. J. Biol. Chem. 279: 25665, 2004. 51. Amadesi, S., Nie, J., Vergnolle, N., Cottrell, G.S., Grady, E.F., Trevisani, M., Manni, C., Geppetti, P., McRoberts, J.A., Ennes, H., Davis, J.B., Mayer, E.A., and Bunnett, N.W. Protease activated receptor 2 sensitizes the capsaicin receptor transient receptor potential vanilloid receptor 1 to induce hyperalgesia. J. Neurosci. 24: 4300, 2004. 52. Dai, Y., Moriyama, T., Higashi, T., Togashi, K., Kobayashi, K., Yamanaka, H., Tominaga, M., and Noguchi, K. Proteinase activated receptor 2-mediated potentiation of transient receptor potential vanilloid subfamily 1 activity reveals a mechanism for proteinaseinduced inﬂammatory pain. J. Neurosci. 24: 4293, 2004. 53. Stander, S., Moorman, C., Schumacher, M., Buddenkotte, J., Artuc, M., Shpacovitch, V., Brzoska, T., Lippert, U., Henz, B.M., Luger, T.A., Metze, D., and Steinhoff, M. Expression of vanilloid receptor subtype-1 in cutaneous sensory nerve ﬁbers, mast cells, and epithelial cells of appendage structure. Exp. Dermatol. 13: 129, 2004. 54. Dattilio, A. and Vizzard, M.A. Up-regulation of protease activated receptors in bladder after cyclophosphamide-induced cystitis and colocalization with capsaicin receptor (VR1) in bladder nerve ﬁbers. J. Urol. 173: 635, 2005. 55. Toth, A., Wang, Y., Kedei, N., Tran, R., Pearce, L.V., Kang, S.U., Jin, M.K., Choi, H.K., Lee, J., and Blumberg, P.M. Different vanilloid agonists cause different patterns of calcium responses in CHO cells heterogenously expressing rat TRPV1. Life Sci. 76: 2921, 2005.

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56. Mohapatra, D.P. and Nau, C. Regulation of Ca2+-dependent desensitization in the vanilloid receptor TRPV1 by calcineurin and cAMP-dependent protein kinase. J. Biol. Chem. 280: 13424, 2005. 57. Mandadi, S., Numazaki, M., Tominaga, M., Bhat, M.B., Armati, P.J., and Roufogalis, B.D. Activation of protein kinase C reverses capsaicin-induced calcium-dependent desensitization of TRPV1 ion channels. Cell Calcium 35: 471, 2004. 58. Chou, M.Z., Mtui, T., Gao, Y.D., Kohler, M., and Middleton, R.E. Resiniferatoxin binds to the capsaicin receptor (TRPV1) near the extracellular side of the S4 transmembrane domain. Biochem. 43: 2501, 2004. 59. Appendino, G. and Szallasi, A. Clinically useful vanilloid receptor TRPV1 antagonists: just around the corner (or too early to tell)? Progress Med. Chem. 44: 145, 2006. 60. Szallasi, A. and DiMarzo, V. New perspectives on enigmatic vanilloid receptors. Trends Neurosci. 23: 491, 2000. 61. Southall, M.D., Li, T., Gharibova, L.S., Pei, Y., Nicol, G.D., and Travers, J.B. Activation of epidermal vanilloid receptor-1 induced release of proinﬂammatory mediators in human keratinocytes. J. Pharmacol. Exp. Ther. 304: 217, 2003. 62. Lazzeri, M., Vannucchi, M.G., Zardo, C., Spinelli, M., Beneforti, P., Turini. D., and Faussone-Pellegrini, M.S. Immunohistochemical evidence of vanilloid receptor 1 in normal human urinary bladder. Eur. Urol. 46: 792, 2004. 63. Golech, S.A., McCarron, R.M., Chen, Y., Bembry, J., Lenz, F., Mechoulam, R., Shohami, E., and Spatz, M. Human brain endothelium: coexpression and function of vanilloid and endocannabinoid receptors. Brain Res. Mol. Brain Res. 132: 87, 2004. 64. Kim, J.C., Beckel, J.M., Birder, L.A., Kiss, S., Washabaugh, C., Kanai, A., Reynolds, I., Dineley, K., de Groat, W.C., and Caterina, M.J. Identiﬁcation of functional vanilloid receptors in human bladder urothelial cells using a nitric oxide microsensor technique and reverse transcriptase polymerase chain reaction. J. Urol. 165: 34, 2001. 65. Dvorakova, M. and Kummer, W. Transient expression of vanilloid receptor subtype 1 in rat cardiomyocytes during development. Histochem. Cell Biol. 116: 223, 2001. 66. Sanchez, J.F., Krause, J.E., and Cortright, D.N. The distribution and regulation of vanilloid receptor VR1 and VR1 5’ splice variant RNA expression in rat. Neurosci. 107: 373, 2001. 67. Reilly, C.A., Taylor, J.L., Lanza, D.L., Carr, B.A., Crouch, D.J., and Yost, G.S. Capsaicinoids cause inﬂammation and epithelial cell death through activation of vanilloid receptors. Toxicol. Sci. 73: 170, 2003. 68. Tian, W., Fu, Y., Wang, D.H., and Cohen, D.M. Regulation of TRPV1 by a novel renally expressed rat TRPV1 splice variant. Am. J. Physiol. Renal Physiol. 290: F117, 2005. 69. Schumacher, M.A., Moff, I., Sudanagunta, S.P., and Levine, J.D. Molecular cloning of an N-terminal domain suggests functional divergence among capsaicin receptor subtypes. J. Biol. Chem. 275: 2756, 2000. 70. Wang, C., Hu, H.Z., Colton, C.K., Wood, J.D., and Zhu, M.X. An alternative splicing product of the murine trpv1 gene dominant negatively modulates the activity of TRPV1 channels. J. Biol. Chem. 279: 37423, 2004. 71. Chen, S.C., Chang, T.J., and Wu, F.S. Competitive inhibition of the capsaicin receptormediated current by dehydroepiandrosterone in rat dorsal root ganglion neurons. J. Pharmacol. Exp. Ther. 311: 529, 2004. 72. Chen, S.C. and Wu, F.S. Mechanisms underlying inhibition of the capsaicin receptormediated current by pregnenolone sulfate in rat dorsal root ganglion neurons. Brain Res. 1027: 196, 2004.

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73. Peroni, R.N., Orliac, M.L., Becu-Villalobos, D., Huidobro-Toro, J.P., Adler-Graschinsky, E., and Celuch, S.M. Sex-linked differences in the vasorelaxant effects of anandamide in vascular mesenteric beds: role of oestrogens. Eur. J. Pharmacol. 493: 151, 2004. 74. Frot, M., Feine, J.S., and Bushnell, M.C. Sex differences in pain perception and anxiety. A psychophysical study with topical capsaicin. Pain 108: 230, 2004. 75. Akiba, Y., Kato, S., Katsube, K., Nakamura, M., Takeuchi, K., Ishii, H., and Hibi, T. Transient receptor potential vanilloid subfamily 1 expressed in pancreatic islet beta cells modulates insulin secretion in rats. Biochem. Biophys. Res. Comm. 321: 219, 2004. 76. Bölcskei, K., Helyes, Zs., Szabó, Á., Sándor, K., Elekes, K., Nèmeth, J., Almási, R., Pintèr, E., Pethö, G., and Szolcsányi, J. Investigation of the role of TRPV1 receptors in acute and chronic nociceptive processes using gene-deﬁcient mice. Pain 117: 368, 2005. 77. Baamonde, A., Lastra, A., Juarez, L., Hidalgo, A., and Menendez, L. TRPV1 desensitization and endogenous vanilloid involvement in the enhanced analgesia induced by capsaicin in inﬂamed tissues. Brain Res. Bull. 67: 476, 2005. 78. Peier, A.M., Moqrich, A., Hergarden, A.C., Reeve, A.J., Andersson, D.A., Story, G.M., Earley, T.J., Dragoni, I., McIntyre, P., Bevan, S., and Patapoutian, A. A TRP channel that senses cold stimuli and menthol. Cell 108: 705, 2002. 79. Bandell, M., Story, G.M., Hwang, S.W., Viswanath, V., Eid, S.R., Petrus, M.J., Earley, T.J., and Patapoutian, A. Noxious cold ion channel TRPA1 is activated by pungent compounds and bradykinin. Neuron 41: 849, 2004. 80. Weil, A., Moore, S.E., Waithe, N.J., Randall, A., and Gunthorpe, M.J. Conservation of functional and pharmacological properties in the distantly related temperature sensors TRPV1 and TRPM8. Mol. Pharmacol. 68: 518, 2005. 81. Sanchez, M.G., Sanchez, A.M., Collado, B., Malagarie-Cazenave, S., Olea, N., Carmena, M.J., Prieto, J.C., and Diaz-Laviada, I.I. Expression of the transient receptor potential vanilloid 1 (TRPV1) in LNCaP and PC-3 prostate cancer cells in human prostate tissue. Eur. J. Pharmacol. 515: 20, 2005. 82. Zhang, L. and Barritt, G.J. Evidence that TRPM8 is an androgen-dependent Ca2+ channel required for the survival of prostate cancer cells. Cancer Res. 64: 8365, 2004. 83. Surh, Y.J. More than spice: capsaicin in hot chili peppers makes tumor cells commit suicide. J. Natl. Cancer Inst. 94: 1263, 2002. 84. Gopinath, P., Wan, E., Holdcroft, A., Facer, P., Davis, J.B., Smith, G.D., Bountra, C., and Anand, P. Increased capsaicin receptor TRPV1 in skin nerve ﬁbers and related vanilloid receptors TRPV3 and TRPV4 in keratinocytes in human breast pain. BMC Women’s Health 5: 2, 2005. 85. Tympanidis, P., Casula, M.A., Yiangou, Y., Terenghi, G., Dowd, P., and Anand, P. Increased vanilloid receptor VR1 innervation of vulvodynia. Eur. J. Pain 8: 129, 2004. 86. Matthews, P.J., Aziz, Q., Facer, P., Davis, J.B., Thompson, D.G., and Anand, P. Increased capsaicin receptor TRPV1 nerve ﬁbers in the inﬂamed human oesophagus. Eur. J. Gastroenterol. Hepatol. 16: 897, 2004. 87. Yiangou, Y., Facer, P., Dyer, N.H., Fowler, C.J., and Anand, P. Vanilloid receptor 1 immunoreactivity in inﬂamed human bowel. Lancet 357: 1338, 2001. 88. Szallasi, A. Vanilloid (capsaicin) receptors in health and disease. Am. J. Clin. Pathol. 118: 110, 2002. 89. Donnerer, J., Liebmann, I., and Schicho, R. Differential regulation of 3-beta-hydroxysteroid dehydrogenase and vanilloid receptor TRPV1 mRNA in sensory neurons by capsaicin and NGF. Pharmacol. 73: 97, 2005.

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90. Carlton, S.M. and Coggeshall, R.E. Peripheral capsaicin receptors increase in the inﬂamed rat hindpaw: a possible mechanism for peripheral sensitization. Neurosci. Lett. 310: 53, 2001. 91. Contassot, E., Tenan, M., Schnuriger, V., Pelte, M.F., and Dietrich, P.Y. Arachidonyl ethanolamine induces apoptosis of uterine cervix cancer cells via aberrantly expressed vanilloid receptor-1. Gynecol. Oncol. 93: 182, 2004. 92. Johansen, M.E., Reilly, C.A., and Yost, G.S. TRPV1 antagonists elevate cell surface populations of receptor protein and exacerbate TRPV1-mediated toxicities in human lung epithelial cells. Tox. Sci. 89: 278, 2006. 93. Doherty, M.J., Mister, R., Pearson, M.G., and Calverley, P.M.C. Capsaicin responsiveness and cough in asthma and chronic obstructive pulmonary disease. Thorax 55: 643, 2000. 94. Mitchell, J.E., Campbell, A.P., New, N.E., Sadofsky, L.R., Kastelik, J.A., Mulrennan, S.A., Compton, S.J., and Morice, A.H. Expression and characterization of the intracellular vanilloid receptor (TRPV1) in bronchi from patients with chronic cough. Exp. Lung Res. 31: 295, 2005. 95. Moqrich, A., Hwang, S.W., Earley, T.J., Petrus, M.J., Murray, A.N., Spencer, K.S., Andahazy, M., Stoey, G.M., and Patapoutian, A. Impaired thermosensation in mice lacking TRPV3, a heat and camphor sensor in the skin. Science 307: 1468, 2005. 96. Vaishnava, P. and Wang, D.H. Capsaicin sensitive sensory nerves and blood pressure regulation. Curr. Med. Chem. Cardiovasc. Hematol. Agents 1: 177, 2003.

7

TRPV2: A CalciumPermeable Cation Channel Regulated by Insulin-Like Growth Factors Itaru Kojima and Masahiro Nagasawa Gunma University

CONTENTS Requirement of Calcium Entry for Cell-Cycle Progression.................................105 Property of the IGF-Regulated Channel ...............................................................106 Molecular Identiﬁcation of the IGF-Regulated Channel ......................................107 Regulation of TRPV2 by IGF-I ............................................................................107 Regulation of Translocation of TRPV2 ................................................................109 References..............................................................................................................112

REQUIREMENT OF CALCIUM ENTRY FOR CELL-CYCLE PROGRESSION Actions of growth factors are essential for mammalian cells to proliferate. For example, ﬁbroblasts continue to grow in the presence of growth factors in serum, and the removal of serum attenuates proliferation. Serum-deprived cells eventually leave the cell cycle, fall into the G0 state, and become quiescent. Quiescent cells reenter the cell cycle when exposed to serum and progress toward the S phase. Two classes of growth factors exist in serum: the competence factor and the progression factor.1 The competence factor activates the quiescent cells, forces them to enter the cell cycle again, and renders them competent to progress through the G1 phase. Then the progression factor acts and brings the competent cells toward the S phase. Hence, it is the progression factor that promotes the cells to progress through the G1 phase to the S phase. A major competence factor in serum is platelet-derived growth factor (PDGF), while a major progression factor in serum is insulin-like growth factor-I (IGF-I).2 The competence factor exerts its action by acting transiently, whereas the progression factor should act continuously: when the progression factor is removed during the G1 phase, the cell-cycle progression is blocked immediately. When the

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progression factor is restored within three hours, cells again progress to the S phase upon readdition of the factor. In contrast, when cells are deprived of the factor for more than three hours, they eventually return to the quiescent state.3 An interesting aspect of the action of the progression factor is that extracellular calcium is absolutely necessary to promote cell-cycle progression.3 When extracellular calcium concentration is reduced to less than 0.3 mM, IGF-I is not able to exert its action as a progression factor.3 An inorganic calcium channel blocker—for example, cobalt or nickel—also blocks the action of IGF-I on cell-cycle progression. Interestingly, when calcium entry is blocked for more than three hours during the G1 phase, cells return to the quiescent state even in the presence of IGF-I. Attenuation of calcium entry is equivalent to the removal of the growth factor. These observations suggest that IGF-I stimulates calcium entry, which is the prerequisite for cell-cycle progression. In accordance with this notion, IGF-I increases the calcium inﬂux rate in competent ﬁbroblasts, and this effect lasts as long as IGF-I is present.4 It is well known that the IGF-I receptor resembles the insulin receptor and has an intrinsic tyrosine kinase activity. Binding of IGF-I to the receptor leads to phosphorylation of many substrates including insulin receptor substrates (IRSs). Phosphorylated IRSs act as docking proteins and eventually activate the Ras and phosphatidylinositol (PI) 3-kinase pathways.5 In addition to the activation of the tyrosine phosphorylation cascade, the IGF-I receptor also continuously activates the calcium entry pathway. Tyrosine phosphorylation of IRSs and subsequent activation of the Ras and PI 3-kinase pathway are not affected by the removal of extracellular calcium or an addition of cobalt or nickel. In this regard, activation of PI 3-kinase and the Ras pathway is independent of the calcium inﬂux pathway. Transfection of the dominant negative Ras does not affect the IGF-induced calcium entry, whereas inhibitors of PI 3-kinase inhibit calcium entry.

PROPERTY OF THE IGF-REGULATED CHANNEL An electrophysiological study reveals that IGF-I activates a calcium-permeable channel in ﬁbroblasts. The IGF-regulated channel is a nonselective calciumpermeable cation channel, the activity of which is regulated by the IGF-I receptor.4,6 Interestingly, activation of the channel by IGF-I is not immediate and requires several minutes for full activation. Once activated, however, the calcium-permeable channel remains activated as long as the ligand binds to the receptor, which is gradually inactivated soon after removal of the ligand.6 These properties of the IGF-regulated channel are suitable for regulation of long-term action (i.e., cell growth). We screened the compound that blocks the IGF-regulated cation channel and found that tranilast, an anti-allergic compound known to inhibit calcium entry in mast cells, blocks the IGF-regulated calcium-permeable channel. Indeed, this compound effectively blocks the growth-promoting action of IGF-I in ﬁbroblasts without affecting either PI 3kinase or the Ras activity. Tranilast is also effective in other types of normal cells as well as in cancer cells.7 This raises the possibility that the IGF-regulated calciumpermeable channel can be a molecular target to treat certain types of cancer. The IGF-regulated channel is a ligand-operated voltage-independent cation channel.6 Of interest is the fact that pretreatment of the cells with pertussis toxin completely blocks IGF-induced calcium entry.4 Pertussis toxin-sensitive trimetric

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G protein may be involved in regulating the IGF-regulated calcium-permeable channel.

MOLECULAR IDENTIFICATION OF THE IGF-REGULATED CHANNEL We were interested in the molecular nature of the IGF-regulated channel and identiﬁed it to be TRPV2.8 It is structurally related to the vanilloid receptor channel VR19 and is a mouse homologue of the VR1-like channel, VRL-1.10 Mouse TRPV2 is composed of 756 amino acids with a relative molecular weight of 86,000 dalton. Like other members of the TRP family channels, TRPV2 has six hydrophobic putative transmembrane domains and an additional short hydrophobic stretch between the ﬁfth and sixth hydrophobic domains. The amino terminal segment contains ankyrin-repeat domains. TRPV2 has 40 percent overall amino acid identity with rat VR1 and 11–12 percent with other TRP family channels. There is a potential N-linked glycosylation site between the ﬁfth and sixth transmembrane domains. There are multiple potential phosphorylation sites for protein kinase A (residues 98 and 326), protein kinase C (residues 97, 111, 322, and 734), and tyrosine kinase (residues 106, 223, and 330). When the expression of TRPV2 is examined by Northern blotting, the transcript is abundantly found in the brain, lungs, and liver. RT-PCR analysis reveals that TRPV2 is also expressed in various other tissues and organs, including the gastrointestinal tract, pancreas, kidney, heart, blood vessels, skeletal muscle, and fat tissues. Histologically, the expression of TRPV2 is abundant in neurons, including Purkinje cells, neuroendocrine cells in the intestine and pancreas, and macrophages in the lung and spleen.11

REGULATION OF TRPV2 BY IGF-I As with other members of the TRP family channels, TRPV2 functions as a calciumpermeable cation channel. Unlike other members of the TRPV family, however, TRPV2 has an intriguing property in which localization of the channel protein is regulated by ligands. Indeed, TRPV2 translocates from an intracellular compartment to the plasma membrane in response to IGF-I.8 This is the ﬁrst example of a channel whose trafﬁcking is regulated by extracellular signals. To directly monitor the localization of the channel in living cells, green ﬂuorescent protein (GFP)–tagged TRPV2 (TRPV2-GFP) is expressed in ﬁbroblasts. In a quiescent condition, TRPV2-GFP is barely detected in the plasma membrane (Figure 7.1A). Instead, TRPV2-GFP is distributed diffusely in cytoplasm. Regarding the intracellular localization, the TRPV2-GFP signal colocalizes with the marker of the endoplasmic reticulum but not with that of mitochondria, the trans-Golgi network, endosomes, or lysosomes. Immunoelectron microscopy reveals that TRPV2 is located in the endoplasmic reticulum in unstimulated cells. In cells stimulated with IGF-I for 15 minutes, the TRPV2 signal found in the intracellular compartment is reduced, and some of the TRPV2 becomes detectable in the plasma membrane (Figure 7.1B). Because localization of a membrane-spanning TRPV2 protein is changed by IGF-I action, it

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FIGURE 7.1 Effect of IGF-I on the distribution of TRPV2. CHO cells expressing TRPV2GFP were incubated for ﬁfteen minutes with or without 1 nM IGF-I and the GFP ﬂuorescence was monitored. A: none, B: IGF-I.

seems likely that TRPV2 moves from the endoplasmic reticulum to the plasma membrane carried on the vesicles. However, the vesicles containing TRPV2 have not been detected to date by immunoelectron microscopy. An alternate possibility is that a portion of the endoplasmic reticulum is directly connected to the plasma membrane and transfers the TRPV2 to the plasma membrane. At present, it is not certain whether or not translocation of the TRPV2 channel involves vesicle trafﬁcking. We were also able to monitor translocation of TRPV2 by measuring the current through TRPV2. Figure 7.2A depicts changes in the whole-cell Cs+ current in

FIGURE 7.2 Time course of the effect of IGF-I on the Cs+ current. (A) CHO cells expressing TRPV2 were stimulated for 30 minutes with 1 nM IGF-I, and the changes in the whole-cell Cs+ current were monitored by the perforate mode patch clump. Values are the mean ± S. E. for three experiments. (B) CHO cells expressing TRPV2 were incubated for 30 minutes with 50 μM Dk(62-85) peptide in the presence and absence of 1 nM IGF-I. Changes in the wholecell Cs+ current were monitored. Values are the mean ± S. E. for three experiments. 䊊: without IGF-I, 䊉: with IGF-I.

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ﬁbroblasts stimulated with IGF-I. In ﬁbroblasts, the expression of other members of the TRPV family is negligible by RT-PCR. Since the Cs+ current observed in ﬁbroblasts is inhibited by ruthenium red, the Cs+ current is mostly through the TRPV2 channel. Adding IGF-I induces a gradual increase in the Cs+ current, which reaches the plateau level within twenty minutes. Upon removal of IGF-I, the Cs+ current decreases gradually and returns to the basal level after 60 minutes. Although we cannot rule out the possibility that IGF-I directly modiﬁes the gating of TRPV2, augmentation of the Cs+ current is slow and perhaps largely due to translocation of TRPV2 to the plasma membrane.8

REGULATION OF TRANSLOCATION OF TRPV2 Glucose transporter 4 (GLUT4) is an insulin-regulated glucose transporter, which translocates from an intracellular pool to the plasma membrane in adipocytes and skeletal muscle cells. GLUT4 translocates from the intracellular storage pool to the plasma membrane by moving on microvesicles.12 Under basal conditions, some small portion of the GLUT4-containing vesicles moves to the plasma membrane. Simultaneously, some of the GLUT4 undergoes endocytosis and eventually returns to the intracellular pool. Therefore, even in an unstimulated condition, GLUT4 in the plasma membrane is in a dynamic equilibrium of exocytosis and endocytosis. When stimulated with insulin, a large amount of GLUT4-containing vesicles leaves the storage pool and moves toward the plasma membrane. This results in a large increase in the amount of GLUT4 expressed in the plasma membrane. Concomitantly, GLUT4 undergoes endocytosis and returns to the intracellular pool. Collectively, a large portion of GLUT4 recycles in insulin-stimulated conditions. We examined whether trafﬁcking of TRPV2 resembles that of GLUT4 by monitoring the Cs+ current in ﬁbroblasts. Under basal conditions, the Cs+ current is low. We ﬁrst addressed whether or not TRPV2 shuttles under basal conditions. If so, it would be expected that blocking internalization of TRPV2 would lead to an increase in the amount of TRPV2 in the plasma membrane and increase the Cs+ current. To examine this experimentally, we needed to block internalization of TRPV2 from the plasma membrane. To this end, we used Dk(62–85), a synthetic peptide derived from the α1 domain of the murine major histocompatibility complex class I antigen known to inhibit endocytosis of GLUT4 and transferrin.13 When Dk(62–85) is added to an unstimulated ﬁbroblast, the Cs+ current increases slightly (Figure 7.2B). Upon removal of the Dk(62–85), the Cs+ current decreases and returns to the basal value. Therefore, a small fraction of TRPV2 shuttles even under basal conditions. Presumably, TRPV2 undergoes endocytosis by a mechanism similar to those of GLUT4 and transferrin. In fact, transfection of the dominantly negative mutant of dynamin, a GTP-binding protein involved in endocytosis of various membrane proteins, including GLUT414 blocks endocytosis of TRPV2. TRPV2 undergoes endocytosis by a mechanism involving dynamin GTPase. The second issue is whether or not the major site of action of IGF-I stimulates exocytotic recruitment of TRPV2 to the plasma membrane. To examine this, we again used Dk(62–85) to block endocytosis. Using Dk(62–85), we were able to assess the unidirectional exocytotic recruitment of TRPV2. Indeed, an addition of IGF-I in Dk(62–85)-treated ﬁbroblasts resulted in a large increase in

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the Cs+ current (Figure 7.2B). Since endocytosis of TRPV2 is blocked in these cells, the results demonstrate that the major site of action of IGF-I on TRPV2 trafﬁcking is the stimulation of the exocytotic step of TRPV2 (Figure 7.2B). Collectively, most of the TRPV2 is located in the endoplasmic reticulum in unstimulated ﬁbroblasts. Some small portion of TRPV2 moves to the plasma membrane, which is balanced by the endocytosis of TRPV2 (Figure 7.3). When cells are stimulated with IGF-I, a relatively large amount of TRPV2 is recruited from the intracellular pool and translocates to the plasma membrane. As a result, the expression of TRPV2 in the plasma membrane increases considerably and calcium entry is augmented. Some portion of TRPV2 also is internalized by endocytosis. When the action of IGF-I is removed, the supply of TRPV2 from the intracellular pool is terminated. In addition, TRPV2 expressed in the plasma membrane undergoes endocytosis and returns to the endoplasmic reticulum. Consequently, the amount of TRPV2 expressed in the plasma membrane is reduced gradually (Figure 7.3). Thus, the major site of action of IGF-I is the recruitment of TRPV2 to the plasma membrane. IGF-I-induced translocation of TRPV2 is blocked by inhibitors of PI 3-kinase, LY294002, and wortmannin. Translocation is also blocked by transfection of the dominant-negative mutant of the p85 subunit of the PI 3-kinase. Although activation of PI 3-kinase is required for IGF-I-induced translocation of TRPV2, it is not a sufﬁcient signal. Thus, activation of PI 3-kinase by adding phosphatidylinositol 3,4,5-trisphosphate does not induce translocation. Obviously, an additional signal is necessary to mobilize TRPV2 from the intracellular compartment. At present, little is known about the molecular mechanism downstream of the PI 3kinase. In this regard, an interesting machinery regulated by PI 3-kinase is the cytoskeletal proteins involved in cell motility. When actin ﬁlaments are disrupted by latrunculin A or cytochalasin D, distribution of TRPV2 is altered considerably. Under basal conditions, most of the TRPV2-GFP localizes in the endoplasmic

FIGURE 7.3 Regulation of TRPV2 by IGF-I. (A) In unstimulated conditions, most of TRPV2 locates in the storage pool (endoplasmic reticulum), and a small fraction of TRPV2 is recycling. (B) When IGF-I is added, TRPV2 is recruited from the storage pool and translocates to the plasma membrane. Simultaneously, TRPV2 is internalized and recycled.

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FIGURE 7.4 Effect of latrunculin A on IGF-I-induced Cs+ current. Cells were preincubated with or without 50μM latrunculin A and then 1 nM IGF-I was added. Changes in the Cs+ current were monitored. 䊊: without latrunculin A, 䊉: with latrunculin A.

reticulum. However, TRPV2 does not translocate to the plasma membrane after the stimulation with IGF-I. Similarly, the Cs+ current is only slightly higher in the basal condition but does not respond to IGF-I when the actin ﬁlament is disrupted by latrunculin A or cytochalasin D (Figure 7.4). Disruption of the actin ﬁlament does not affect the decrease in the TRPV2 current, suggesting that the actin ﬁlament is not necessary for endocytosis of TRPV2. These results indicate that the actin ﬁlament plays a critical role in recruitment of TRPV2 to the plasma membrane from the endoplasmic reticulum. In contrast, disruption of microtubules by nocodazole does not affect the translocation of TRPV2 induced by IGF-I. Among various cytoskeletal proteins, reorganization of the actin ﬁlament may be involved in IGF-I–mediated translocation of TRPV2. Many issues still remain unsolved. First, the molecular mechanism regulating translocation is largely elusive. For example, it is not totally certain whether or not vesicle trafﬁcking is involved in translocating TRPV2. Suppose TRPV2 is carried on vesicles, the formation, trafﬁcking, and exocytosis of the vesicles should be regulated by the IGF-I signal. These regulatory mechanisms need to be identiﬁed. Second, it is not clear at present that translocation is the sole mechanism by which IGF-I regulates TRPV2. In other words, it is possible that IGF-I also modulates TRPV2 gating. If this is the case, it is necessary to identify the regulatory mechanism. Third, TRPV2 is located in the endoplasmic reticulum under basal conditions. It is not certain that TRPV2 functions as a calcium-permeable channel in this organella.

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If so, trafﬁcking of TRPV2 may alter the calcium handling in the endoplasmic reticulum. Fourth, translocation of TRPV2 is regulated by IGF-I in ﬁbroblasts and neuroendocrine cells,8 but other ligands may regulate TRPV2 in other types of cells. In fact, translocation of TRPV2 is induced by ligands that bind to G-protein-coupled receptors in other types of cells.15 Regulation of the TRPV2 translocation by those ligands may be different. It is also shown that membrane stretch induces translocation of TRPV2 in myocytes.16 Translocation may be a major regulatory mechanism for TRPV2 activation. Further works are clearly required to elucidate the mechanism and the role of TRPV2 translocation.

REFERENCES 1. Scher, C.D. et al. Platelet-derived growth factor and the regulation of the mammalian ﬁbroblast cell cycle. Biochem. Biophys. Acta 560, 217, 1979. 2. Stiles, C.D. et al. Dual control of cell growth by somatomedins and platelet-derived growth factor. Proc. Natl. Acad. Sci. USA 76, 1279, 1979. 3. Kojima, I. et al. Role of calcium entry and protein kinase C in the progression activity of insulin-like growth factor-I in Balb/c 3T3 cells. J. Biol. Chem. 268, 10003, 1993. 4. Kojima, I. et al. Calcium inﬂux: an intracellular message of the mitogenic action of insulin-like growth factor-I. J. Biol. Chem. 263, 16561, 1988. 5. Nakae, J., Kido, Y., and Accili, D. Distinct and overlapping function of insulin and IGF-I receptors. Endocrine Rev. 22, 818, 2001. 6. Matsunaga, H. et al. Activation of a calcium-permeable cation channel by insulinlike growth factor-Ii in Balb/c 3T3 cells. Am. J. Physiol. 255, C442, 1988. 7. Nie, L. et al. Inhibition of proliferation of MCF-7 breast cancer cells by a blocker of Ca2+-permeable channel. Cell Calcium 22, 75, 1997. 8. Kanzaki, M. et al. Translocation of a calcium-permeable channel induced by insulinlike growth factor-I. Nature Cell Biol. 1, 165, 1999. 9. Caterina, M.J. et al. The capsaicin receptor: a heat-activated ion channel in the pain pathway. Nature 389, 816, 1997. 10. Caterina, M.J. et al. A capsaicin receptor homologue with a high threshold for noxious heat. Nature 398, 436, 1999. 11. Kowase, T. et al. Immunohistochemical localization of growth factor–regulated channel (GRC) in human tissues. Endocrine J. 49, 349, 2002. 12. Haney, P.M. et al. Intracellular targeting of the insulin-regulatable glucose transporter (GLUT4) is isoform speciﬁc and independent of cell type. J. Cell Biol. 114, 689, 1991. 13. Shibata, H. et al. Dissection of GLUT4 recycling pathway into exocytosis and endocytosis in rat adipocytes. J. Biol. Chem. 270, 11489, 1995. 14. Omata, W. et al. Subcellular distribution of GLUT4 in CHO cells overexpressing mutant dynamin. Biochem. Biophys. Res. Commun. 241, 401, 1997. 15. Boels, K. et al. The neuropeptide head activator induces activation and translocation of the growth factor–regulated Ca2+-permeable channel GRC. J. Cell Sci. 114, 3599, 2001. 16. Iwata, Y. et al. A novel mechanism of myocyte degeneration involving the Ca2+permeable growth factor–regulated channel. J. Cell Biol. 161, 957, 2003.

8

Molecular Mechanisms of TRPV4 Gating Stefan Heller Stanford University School of Medicine

Roger G. O’Neil The University of Texas Health Science Center

CONTENTS Introduction............................................................................................................113 TRPV4 Structure ...................................................................................................114 Amino Terminus ........................................................................................114 Membrane-Spanning Core Region and Pore Loop...................................115 Carboxyl Terminus ....................................................................................116 Emerging Mechanism of Channel Activation .......................................................116 Mechanosensitivity ....................................................................................116 Thermosensitivity ......................................................................................117 Diacylglycerol and Phorbol Esters............................................................118 Arachidonic Acid and Epoxygenase Metabolites .....................................119 Molecular Insights into Channel Selectivity and Gating......................................119 The Molecular Basis of Selectivity...........................................................119 Emerging Paradigms of Gating .................................................................120 Summary ................................................................................................................120 References..............................................................................................................121

INTRODUCTION TRPV4, the fourth member of the vanilloid subfamily of TRP channels, is a calciumpermeable cation channel that is detectable in both sensory and nonsensory cells. It has been shown to be widely expressed, including in kidneys, lungs, hearts, brains, endothelial cells, and dorsal root and trigeminal sensory ganglia. The channel is characterized by multimodal activation properties that implicate it in a broad range of functions from osmoregulation to thermosensing. Although TRPV4 was originally identiﬁed as an osmotically activated channel,1–3 recent evidence demonstrates that the channel can be activated by diverse stimuli including hypoosmotic swelling,1–3 shear stress,4 nonnoxious temperatures,5,6 acidity,7 phorbol esters (both protein 113

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kinase C–activating and nonactivating phorbol esters),4,8,9 and downstream metabolites of arachidonic acid (epoxyeicosatrienoic acids).10,11 The mechanism(s) underlying TRPV4 activation by these differing modalities remain(s) poorly understood. However, new evidence points to some common pathways of activation by these diverse stimuli that may underlie the apparent broad range of functions associated with the channel.

TRPV4 STRUCTURE The 871 amino acids (aa) that make up the most common isoform of TRPV4 are predicted to form the individual subunits of the functional homotetrameric ion channel. Amino and carboxyl termini of TRPV4 are localized in the cytoplasm and ﬂank the core region consisting of six transmembrane-spanning domains (TM1–6) and a pore loop between TM5 and TM6.1–3 Determination of the structure of the bacterial P-loop KcsA potassium channel,12,13 as well as other recently determined structures for KirBac1.114 and KirBac3.1,15 has provided new insights into the general structure of P-loop channels, prompting speculation as to the potential structure of TRPV channels.16 P-loop channels typically form as tetramers, consistent with TRP channel structure, with a four-fold symmetry around the central pore region.17,18

AMINO TERMINUS More than half of the TRPV4 protein is formed by the intracellularly located amino terminus that most prominently harbors three ankyrin repeat domains (ARD1–3) within an ankyrin repeat region from aa235 to aa367, a cluster of four protein kinase C (PKC)–phosphorylation sites and a cAMP-dependent–phosphorylation site upstream of the ankyrin repeat region, and a cluster of two PKC sites within and downstream of ARD3 (Figure 8.1A). A functionally identiﬁed Src family dependent tyrosine phosphorylation site has been described at aa253 within ARD1. Other potentially functionally important motifs are an N-myristoylation site at aa24–29, a bipartite nuclear targeting sequence that overlaps with part of ARD3, and a PKC phosphorylation site. It has been hypothesized that increased PKC activity in response to mechanical stress could be a factor in TRPV4 activation. Activation of PKC with phorbol esters leads to opening of TRPV4 and increase of intracellular Ca2+ concentration when the experiment is conducted at 37°C.4,9 At room temperature, PKCactivating phorbol esters have no or very little effect on TRPV4 gating. The sites of PKC action on TRPV4 are unknown. Candidates for direct phosphorylation are the six PKC phosphorylation sites in the amino terminus and one motif just downstream of TM2 (Figure 8.1A–B). Another motif that has been implicated in regulating TRPV4 is a single Src family tyrosine phosphorylation site within ARD1 that is phosphorylated in response to hypotonic stress.19 A Y253F point mutant exhibited nearly abolished hypotonicity responses,19 a result that recently has been challenged.20,21

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FIGURE 8.1 Schematic overview of TRPV4’s predicted structural and functional components. Shown are schematic representations of TRPV4’s amino terminus (A), its central region with the membrane-spanning domains and the pore loop (B), and the channel’s carboxyl terminus (C). Speciﬁc domains and amino acids are indicated. Regions that are predicted to be extracellularly located are indicated with a black horizontal bar in (B). Also shown in (B) are the proposed pore helix and selectivity ﬁlter displaying the TIGMGD region similar to the K+ channel selectivity ﬁlter signature sequence.

MEMBRANE-SPANNING CORE REGION

AND

PORE LOOP

The 242-aa-long central domain of TRPV4 consists of TM1–6, which feature, between TM5 and TM6, a short hydrophobic stretch that is the putative pore region or pore loop (PL, Figure 8.1B). TRPV4 does display only a few predicted short extracellular domains, 25 aa between TM1 and TM2, 4 aa between TM3 and TM4, and 34 aa and 12 aa upstream and downstream of the PL. The channel appears to be posttranslationally modiﬁed by glycosylation,22 and a bona ﬁde Asn glycosylation site within the extracellular stretch between TM5 and the PL has been shown to be glycosylated in heterologously expressed TRPV4.23 A PKC phosphorylation site downstream of TM2 is potentially involved in the above-mentioned PKC regulation of TRPV4 activation. By analogy to the bacterial KcsA K+ channel,12,13 the PL is probably composed of a short highly hydrophobic pore helix immediately followed by an ion selectivity segment, the channel selectivity ﬁlter, that has been postulated to encompass aa672–682.16 Several individual aas affect the selectivity and permeability; when mutated (notably aspartate residues D672 and D682, and, somewhat surprisingly,

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M680), they markedly impair calcium permeation, results are consistent with the purported selectivity ﬁlter of TRPV4 (see below).

CARBOXYL TERMINUS TRPV4’s carboxyl terminal tail appears to be the docking site for at least two interacting proteins. One of these sites has been narrowed down to a region between aa812 and aa831 and confers binding to calmodulin.24 Mutations within this region resulted in a loss of Ca2+-dependent calmodulin binding and a loss of Ca2+-dependent potentiation of TRPV4 currents. This observed effect adds to the complex Ca2+ dependence of TRPV4 activity that also involves inhibitory effects.1,25 Immediately upstream of the calmodulin-binding region, an interaction site with microﬁlament-associated protein 7 (MAP7) has been proposed. Coexpression of TRPV4 with MAP7 in CHO cells apparently increases the amount of TRPV4 protein associated with the plasma membrane, which could be a method employed by cells to control the density of TRPV4 in the plasma membrane.26

EMERGING MECHANISM OF CHANNEL ACTIVATION TRPV4 is a multimodal channel regulated by a diverse array of stimuli, implicating multiple mechanisms of regulation and function. The potential cellular pathways underlying this regulation are outlined below.

MECHANOSENSITIVITY The TRPV4 channel was initially cloned based on its sensitivity to hypoosmotic cell swelling, implicating the channel as a potential mechanosensitive channel1–3 as mentioned. It was subsequently shown that application of ﬂuid shear stress (or ﬂuid ﬂow) across the apical surface of TRPV4-expressing HEK293 epithelial cells and M-1 renal collecting duct cells activated the channel, conﬁrming the mechanosensitive nature of activation.4,27 The response to mechanical stimulation, however, was shown to be highly sensitive to temperature,1,4 implicating a temperature-induced augmentation or sensitization to mechanical stimuli. The mechanosensitive nature of TRPV4, or a TRPV4 complex, was most clearly demonstrated in a seminal study of C. elegans using osm-9 mutant worms lacking expression of a worm TRPV homologue, OSM-9, which underlies osmotic (hyperosmolarity) and mechanical (nose touch) avoidance behavior. Transgenically targeting expression of mammalian TRPV4 to the ASH amphid sensory neuron in the mutants (a normal site of OSM-9 expression) rescued the osmotic and mechanical avoidance responses, providing compelling evidence that in the in vivo setting, TRPV4 functions as an osmosensitive and mechanosensitive channel or sensory complex.28 The mechanism by which the TRPV4 channel is activated by mechanical stresses is currently not fully understood. Two, not mutually exclusive, principal mechanisms have been proposed: direct mechanical activation and indirect activation through other transduction pathways. Cell swelling can lead to activation of both the phospholipase C (PLC)/diacylglycerol (DAG) transduction pathway29,30

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and the phospholipase A2 (PLA2)/arachidonic acid (AA) transduction pathway,31–33 two pathways that can modulate, or regulate, TRPV4 activation (see below). However, Nilius and coworkers demonstrated, in a detailed series of studies, that swelling-induced production of AA and its downstream epoxyeicosatrienoic acid (EET) metabolites (5,6-EET and 8,9-EET) may be responsible, at least in part, for hypotonic-induced activation of TRPV4.10,11,21 Inhibition of PLA2 or cytochrome P450 epoxygenase activity reduced the swelling-induced activation of TRPV4, while application of the EET metabolites or inhibition of EET hydrolysis enhanced both the swelling and AA-induced activation of TRPV4. TRPV4 knockout mice demonstrated a reduced swelling and AA-induced activation of calcium entry in mouse aortic endothelial cells.10 Hence, the PLA2–cytochrome P450 pathway may reﬂect a dominant pathway and mechanism by which mechanical stress may activate the channel. However, direct application of membrane stretch per se on channel activation, such as recently demonstrated for TRPC1,34 without input from arachidonic acid and its metabolites, or from DAG and its metabolites, has not been systematically tested and remains to be determined, particularly at physiologically relevant temperatures. The mechanosensory functions of TRPV4 in mammals are slowly being elucidated as described in more detail previously35,36 and in other chapters of this volume. As noted, TRPV4 has been shown to be activated by cell swelling,1–3 implicating a potential role in cell volume regulation,37 and in sensing shear stress, where the channel may play a key sensory role in ﬂow-sensitive tissues such as vascular endothelial cells and renal tubular epithelial cells.4,6,27 The channel has been strongly implicated as a mechanosensor in cilia of oviductal epithelial cells, showing an increased channel activity and accompanying Ca2+ inﬂux, following an increase in ﬂuid viscosity (i.e., increased mechanical load).38 In addition, hypotonicityinduced nociception has also been described as TRPV4-mediated.39 Further, studies employing TRPV4-deﬁcient knockout mice have implicated TRPV4 in speciﬁc functions: in sensing plasma osmolarity by the sensory circumventricular organs of the hypothalamus, which, in turn, regulate secretion of antidiuretic hormone and control of extracellular ﬂuid osmolarity;40,41 in sensing tail pressure by dorsal root ganglia sensory neurons, implicating TRPV4 as a high threshold mechanoreceptor;7 and in sensing hypotonic cell volume in mouse aortic endothelial cells.10 Other functions of TRPV4 are continuing to emerge, and many more are likely to be forthcoming.35,36

THERMOSENSITIVITY The survival of animals, especially warm-blooded animals, relies on the animals’ abilities to sense and respond to changes in temperature. TRP channels may be central components of this sensing ability. The TRPV channel members, TRPV1–4, have been shown to be activated by warm to noxious temperatures and have been implicated in temperature sensing by the body. The TRPV channels display deﬁned temperature thresholds for activation: TRPV1, 42°C;42,43 TRPV2, 52°C;44 TRPV3, 31°C;45–47 and TRPV4, near 27–35°C.5,6 The channels have recently been dubbed “thermoTRPs” because of this feature.48 These investigators demonstrated that the

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mechanism of temperature dependency may relate to a temperature-dependent shift of the channels’ voltage dependency (normally not a notable feature of TRP channels) to more physiologically relevant voltage ranges. That is, elevating the temperature for warm/hot TRPVs provides a left shift in the voltage sensitivity, making the voltage dependency apparent at more “physiological” voltages, thereby activating the channel. (Cool/cold-activated TRPs displayed a complementary right shift in the voltage sensitivity with cooling.) TRPV4, however, does not speciﬁcally ﬁt this model, as it could not be demonstrated to be temperature sensitive when studied in patch-clamp studies where the membrane was detached from the cell.48 Other cellular factors, yet to be identiﬁed, may be critical for the TRPV4 channel to display temperature-induced activation. The physiological function of TRPV4 in thermosensation is still emerging.48,49 Besides functioning as a general sensor of altered cell/tissue temperature, TRPV4 expression in skin keratinocytes has been implicated as a sensor of “warm” temperatures.50 TRPV4 knockout studies clearly implicate TRPV4 in selecting “preferred” footpad temperatures and as being essential in thermal hyperalgesia.51

DIACYLGLYCEROL

AND

PHORBOL ESTERS

The ﬁrst ligand discovered to activate TRPV4 was the synthetic phorbol ester, 4α-phorbol 12,13-didecanoate (4α-PDD).8 It was observed that 4α-PDD could potently activate TRPV4, even at relatively modest concentrations (<1 μM), and it appeared to be relatively speciﬁc for TRPV4 because it does not appear to activate other TRPV family members. Since 4α-PDD is a non-protein kinase C–activating phorphol ester that is not metabolizable, it may activate TRPV4 by directly binding to the channel or an associated subunit. However, recent studies by Xu and coworkers9 have demonstrated that other inactive 4α isomers can activate the channel with a potency that appears to be related to their chemical hydrophobicity. This may indicate a potential effect of the phorbol isomers on the lipid environment in modulating, or activating, the channel. The physiological relevance of such a lipid-modulated mechanism of activation may have broad implications on the function and mechanisms of TRPV4 gating as noted for other mechanogated channels.52,53 The PKC-activating phorbol esters were originally considered only to activate TRPV4 weakly based on studies done at room temperature.8 It was subsequently demonstrated that elevation of temperature to the physiological range resulted in a potent augmentation, or sensitization, of the channel to phorbol esters and mechanical stimuli.4 At 37°C, PMA in the submicromolar range activated TRPV4 in a PKC-dependent manner, approaching the potency of 4α-PDD.4,9 Other PKCactivating phorbol esters (phorbol 12,13-didecanoate, PDD; phorbol 12,13dibutyrate, PDBu)9 similarly activated the channel in a PKC-dependent manner, implicating PKC as an important modulator of TRPV4 activity. These studies raise the exciting possibility that signaling pathways underlying induction of PKC, such as G-protein-coupled receptors or tyrosine kinase receptors linked to activation of phospholipase C, may reﬂect emerging, as yet undiscovered, pathways for TRPV4 activation.

Molecular Mechanisms of TRPV4 Gating

ARACHIDONIC ACID

AND

119

EPOXYGENASE METABOLITES

The discovery that arachidonic acid (AA) and its downstream metabolites may activate TRPV4 provided the ﬁrst evidence for endogenous agonists of the channel. It was initially shown by Nilius and coworkers11 that the endocannabinoid anandamide (AEA) and its lipoxygenase (fatty acid amidohydrolase) metabolite, AA, could activate TRPV4. A nonmetabolizable analogue of AEA (e.g., methanandamide) was without effect, while an inhibitor of fatty acid amidohydrolase, phenylmethylsulfonyl ﬂuoride, abolished the AEA-induced activation of TRPV4. It was subsequently shown that generating EET metabolites of AA via the cytochrome P450 epoxygenase pathway, notably 5,6-EET and 8,9-EET, could activate TRPV4.10,21 Other studies have shown that the 11,12-EET may similarly activate TRPV4.54 Furthermore, activation by AA metabolites appears to be distinct from the 4α-PDD pathway of activation because mutation of a tyrosine residue near the third transmembrane domain (Y555, Figure 8.1B) to an alanine impairs activation by 4α-PDD, but not by AA or cell swelling, implicating a potential role for tyrosine phosphorylation in the activation pathway for 4α-PDD.21 Recent studies in mouse vascular endothelial cells, which express endogenous TRPV4, further demonstrate that inhibiting EET hydrolysis also leads to an enhanced response of AA or EET analogues, while in cells isolated from animals lacking TRPV4 (TRPV4−/−), all TRPV4 agonists are without effect.10 Hence, accumulating evidence points to the AA pathway and the epoxygenase metabolites as playing key roles in regulating TRPV4 activity in the endogenous setting.

MOLECULAR INSIGHTS INTO CHANNEL SELECTIVITY AND GATING The selectivity ﬁlter and gating mechanism of TRPV4 have not been determined. Insights into these mechanisms are beginning to emerge from recent studies of TRPV4 and other P-loop channels.

THE MOLECULAR BASIS

OF

SELECTIVITY

The molecular basis of the TRPV4 channel selectivity ﬁlter has not been fully elucidated, although charged residues in the purported ﬁlter segment, aa672–682, have been implicated. In particular, two aspartate residues (D672 and D682) appear to cooperatively affect the pore’s Ca2+ permeability and channel rectiﬁcation.55 Nevertheless, these two aas are not strictly linked as mutations to neutralize D672 alone does not have any strong effects, and neutralization of D682 but not D672 almost abolishes the blocking effect of ruthenium red.20,55 Nilius and colleagues16,20 have hypothesized that the structural design of the TRPV4 pore is comparable to that of the bacterial KcsA K+ channel.12,13 Based on this assessment, the PL is characterized by a short pore helix followed by the selectivity ﬁlter (Figure 8.1B). This notion was spurred by the weak but noticeable similarity of the TIGMGD stretch of aa677–aa682 (Figure 8.1B) within the PL with the TXGYGD selectivity ﬁlter residues of the K+ channel signature sequence.12,13

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The aforementioned mutations of D672 and D682, as well as M680, which impair Ca2+ permeation and reduce whole cell current amplitude, circumstantially support this hypothesis and clearly demonstrate involvement of this segment of the PL in ion selectivity and permeability.20,55

EMERGING PARADIGMS

OF

GATING

The conformational changes in channel structure that lead to functional transitions from closed to open conducting states (i.e. “gating”) are not well understood for most channels. Again, based on structural analysis of bacterial K+ channels,12–15 potential TRPV4 gating mechanisms can be suggested.16 The tetrameric subunit arrangement provides for fourfold symmetry around the central pore region,17,18 where an inner transmembrane helical segment from each of the four subunits each arranges concentrically to form the channel barrel or pore. The transmembrane helices are tilted inward to form a cone shape of the barrel with narrowing toward the cytoplasmic face. This narrowing of the transmembrane segments toward the cytoplasmic face forms the main activation gate.17,18,56 Hinging movements of these segments can widen the pore, thereby opening the gate, or can narrow the pore, thereby closing the gate. The overall structure of the TRPV channels, including TRPV4, would be consistent with such a gating structure, as speculated by others,16 although this model for TRPV4 gating has not been explicitly evaluated. Other components of the structure of P-loop channels, notably the pore helix, may also play a role in the gating process. Rotation or movement of the pore helix can modulate gating, or perhaps act as a secondary gate, in some P-loop channels.57,58 Recent studies of TRPV5 gating, using the cysteine-accessibility method, have shown that inhibiting the channel by internal hydrogen is associated with a clockwise rotation of the pore helix along its long axis.59 It is implicit in these ﬁndings that such rotational movements of the pore helix may be critical for gating other TRPV channels, such as TRPV4. Therefore, it is reasonable to conclude that, at a minimum, both hinging movements of the helices forming the activation gate at the cytoplasm face and rotation of the pore helices of the pore loop regions are components of TRPV channel gates. For multimodal channels, such as TRPV4, it may be that other structural components to gating may also be at play. Hence, future studies should provide exciting new insights into the molecular mechanisms of gating of TRPV4 and other TRP channels.

SUMMARY In the ﬁve years since TRPV4 was ﬁrst described, substantial progress has been made toward understanding this channel’s roles in cellular and physiologic processes. These studies revealed that TRPV4 is a rather complex ion channel, capable of integrating multiple physical stimuli such as mechanic stress and temperature. In addition, the channel’s gating is considerably modulated by phosphorylation, external and internal Ca2+ concentration, PLC activity, and AA metabolites, which are potentially generated via the PLA2 pathway. The strong activating capacity of certain synthetic non-PKC-activating phorbol esters adds lipid modulation as an intriguing

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additional aspect of TRPV4 activation. Future structure and function studies, not only of TRPV4 but also of other TRP channels, will shed more light on the physiological purpose of the apparently complex gating and modulation of TRPV4.

REFERENCES 1. Liedtke, W., Choe, Y., Marti-Renom, M.A., Bell, A.M., Denis, C.S., Sali, A., Hudspeth, A.J., Friedman, J.M., and Heller, S., Vanilloid receptor–related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor, Cell 103 (3), 525–35, 2000. 2. Strotmann, R., Harteneck, C., Nunnenmacher, K., Schultz, G., and Plant, T.D., OTRPC4, a nonselective cation channel that confers sensitivity to extracellular osmolarity, Nat. Cell Biol. 2 (10), 695–702, 2000. 3. Wissenbach, U., Bodding, M., Freichel, M., and Flockerzi, V., Trp12, a novel Trp-related protein from kidney, FEBS Lett. 485 (2–3), 127–34, 2000. 4. Gao, X., Wu, L., and O’Neil, R.G., Temperature-modulated diversity of TRPV4 channel gating: activation by physical stresses and phorbol ester derivatives through protein kinase C–dependent and –independent pathways, J. Biol. Chem. 278 (29), 27129–37, 2003. 5. Guler, A.D., Lee, H., Iida, T., Shimizu, I., Tominaga, M., and Caterina, M., Heatevoked activation of the ion channel, TRPV4, J. Neurosci. 22 (15), 6408–14, 2002. 6. Watanabe, H., Vriens, J., Suh, S.H., Benham, C.D., Droogmans, G., and Nilius, B., Heat-evoked activation of TRPV4 channels in a HEK293 cell expression system and in native mouse aorta endothelial cells, J. Biol. Chem. 277 (49), 47044–51, 2002. 7. Suzuki, M., Mizuno, A., Kodaira, K., and Imai, M., Impaired pressure sensation in mice lacking TRPV4, J. Biol. Chem. 278 (25), 22664–68, 2003. 8. Watanabe, H., Davis, J.B., Smart, D., Jerman, J.C., Smith, G.D., Hayes, P., Vriens, J., Cairns, W., Wissenbach, U., Prenen, J., Flockerzi, V., Droogmans, G., Benham, C.D., and Nilius, B., Activation of TRPV4 channels (hVRL-2/mTRP12) by phorbol derivatives, J. Biol. Chem. 277 (16), 13569–77, 2002. 9. Xu, F., Satoh, E., and Iijima, T., Protein kinase C–mediated Ca2+ entry in HEK 293 cells transiently expressing human TRPV4, Br. J. Pharmacol. 140 (2), 413–21, 2003. 10. Vriens, J., Owsianik, G., Fisslthaler, B., Suzuki, M., Janssens, A., Voets, T., Morisseau, C., Hammock, B.D., Fleming, I., Busse, R., and Nilius, B., Modulation of the Ca2 permeable cation channel TRPV4 by cytochrome P450 epoxygenases in vascular endothelium, Circ. Res. 97 (9), 908–15, 2005. 11. Watanabe, H., Vriens, J., Prenen, J., Droogmans, G., Voets, T., and Nilius, B., Anandamide and arachidonic acid use epoxyeicosatrienoic acids to activate TRPV4 channels, Nature 424 (6947), 434–38, 2003. 12. Doyle, D.A., Morais Cabral, J., Pfuetzner, R.A., Kuo, A., Gulbis, J.M., Cohen, S.L., Chait, B.T., and MacKinnon, R., The structure of the potassium channel: molecular basis of K+ conduction and selectivity, Science 280 (5360), 69–77, 1998. 13. Zhou, Y., Morais-Cabral, J.H., Kaufman, A., and MacKinnon, R., Chemistry of ion coordination and hydration revealed by a K+ channel-Fab complex at 2.0 A resolution, Nature 414 (6859), 43–48, 2001. 14. Kuo, A., Gulbis, J.M., Antcliff, J.F., Rahman, T., Lowe, E.D., Zimmer, J., Cuthbertson, J., Ashcroft, F.M., Ezaki, T., and Doyle, D.A., Crystal structure of the potassium channel KirBac1.1 in the closed state, Science 300 (5627), 1922–26, 2003.

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15. Kuo, A., Domene, C., Johnson, L.N., Doyle, D.A., and Venien-Bryan, C., Two different conformational states of the KirBac3.1 potassium channel revealed by electron crystallography, Structure (Camb) 13 (10), 1463–72, 2005. 16. Owsianik, G., Talavera, K., Voets, T., and Nilius, B., Permeation and selectivity of TRP channels, Annu. Rev. Physiol., 2005. 17. Doyle, D.A., Structural changes during ion channel gating, Trends Neurosci. 27 (6), 298–302, 2004. 18. MacKinnon, R., Potassium channels, FEBS Lett. 555 (1), 62–65, 2003. 19. Xu, H., Zhao, H., Tian, W., Yoshida, K., Roullet, J.B., and Cohen, D.M., Regulation of a transient receptor potential (TRP) channel by tyrosine phosphorylation. SRC family kinase-dependent tyrosine phosphorylation of TRPV4 on TYR-253 mediates its response to hypotonic stress, J. Biol. Chem. 278 (13), 11520–27, 2003. 20. Nilius, B., Vriens, J., Prenen, J., Droogmans, G., and Voets, T., TRPV4 calcium entry channel: a paradigm for gating diversity, Am. J. Physiol. Cell Physiol. 286 (2), C195–205, 2004. 21. Vriens, J., Watanabe, H., Janssens, A., Droogmans, G., Voets, T., and Nilius, B., Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4, Proc. Natl. Acad. Sci. USA 101 (1), 396–401, 2004. 22. Arniges, M., Fernandez-Fernandez, J.M., Albrecht, N., Schaefer, M., and Valverde, M.A., Human TRPV4 channel splice variants revealed a key role of ankyrin domains in multimerization and trafﬁcking, J. Biol. Chem. 281: 8996, 2006. 23. Xu, H., Fu, Y., Tian, W., and Cohen, D.M., Glycosylation of the osmoresponsive transient receptor potential channel TRPV4 on Asn-651 inﬂuences membrane trafﬁcking, Am. J. Physiol. Renal Physiol. 290: F1103–F1109, 2006. 24. Strotmann, R., Schultz, G., and Plant, T.D., Ca2+-dependent potentiation of the nonselective cation channel TRPV4 is mediated by a C-terminal calmodulin-binding site, J. Biol. Chem. 278 (29), 26541–49, 2003. 25. Watanabe, H., Vriens, J., Janssens, A., Wondergem, R., Droogmans, G., and Nilius, B., Modulation of TRPV4 gating by intra- and extracellular Ca2+, Cell Calcium 33 (5–6), 489–95, 2003. 26. Suzuki, M., Hirao, A., and Mizuno, A., Microtubule-associated [corrected] protein 7 increases the membrane expression of transient receptor potential vanilloid 4 (TRPV4), J. Biol. Chem. 278 (51), 51448–53, 2003. 27. O’Neil, R.G., Wu, L., and Gao, X., Mechanosensitive nature of the TRPV4 channel in renal epithelia revealed by siRNA gene silencing, Faseb J. 19, A1163, 2005. 28. Liedtke, W., Tobin, D.M., Bargmann, C.I., and Friedman, J.M., Mammalian TRPV4 (VR-OAC) directs behavioral responses to osmotic and mechanical stimuli in Caenorhabditis elegans, Proc. Natl. Acad. Sci. USA 100 (Suppl. 2), 14531–36, 2003. 29. O’Neil, R.G. and Leng, L., Osmo-mechanically sensitive phosphatidylinositol signaling regulates a Ca2+ inﬂux channel in renal epithelial cells, Am. J. Physiol. 273 (1 Pt. 2), F120–28, 1997. 30. Suzuki, M., Kawahara, K., Ogawa, A., Morita, T., Kawaguchi, Y., Kurihara, S., and Sakai, O., [Ca2+]i rises via G protein during regulatory volume decrease in rabbit proximal tubule cells, Am. J. Physiol. 258 (3 Pt. 2), F690–96, 1990. 31. Basavappa, S., Pedersen, S.F., Jorgensen, N.K., Ellory, J.C., and Hoffmann, E.K., Swelling-induced arachidonic acid release via the 85-kDa cPLA2 in human neuroblastoma cells, J. Neurophysiol. 79 (3), 1441–49, 1998. 32. Pedersen, S., Lambert, I.H., Thoroed, S.M., and Hoffmann, E.K., Hypotonic cell swelling induces translocation of the alpha isoform of cytosolic phospholipase A2

Molecular Mechanisms of TRPV4 Gating

33.

34.

35. 36. 37. 38.

39.

40. 41. 42.

43. 44.

45.

46.

47.

48.

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but not the gamma isoform in Ehrlich ascites tumor cells, Eur. J. Biochem. 267 (17), 5531–39, 2000. Thoroed, S.M., Lauritzen, L., Lambert, I.H., Hansen, H.S., and Hoffmann, E.K., Cell swelling activates phospholipase A2 in Ehrlich ascites tumor cells, J. Membr. Biol. 160 (1), 47–58, 1997. Maroto, R., Raso, A., Wood, T.G., Kurosky, A., Martinac, B., and Hamill, O.P., TRPC1 forms the stretch-activated cation channel in vertebrate cells, Nat. Cell Biol. 7 (2), 179–85, 2005. Mutai, H. and Heller, S., Vertebrate and invertebrate TRPV-like mechanoreceptors, Cell Calcium 33 (5–6), 471–78, 2003. O’Neil, R.G. and Heller, S., The mechanosensitive nature of TRPV channels, Pﬂügers Arch., 2005. Becker, D., Blase, C., Bereiter-Hahn, J., and Jendrach, M., TRPV4 exhibits a functional role in cell-volume regulation, J. Cell Sci. 118 (Pt. 11), 2435–40, 2005. Andrade, Y.N., Fernandes, J., Vazquez, E., Fernandez-Fernandez, J.M., Arniges, M., Sanchez, T.M., Villalon, M., and Valverde, M.A., TRPV4 channel is involved in the coupling of ﬂuid viscosity changes to epithelial ciliary activity, J. Cell Biol. 168 (6), 869–74, 2005. Alessandri-Haber, N., Yeh, J.J., Boyd, A.E., Parada, C.A., Chen, X., Reichling, D.B., and Levine, J.D., Hypotonicity induces TRPV4-mediated nociception in rat, Neuron 39 (3), 497–511, 2003. Liedtke, W. and Friedman, J.M., Abnormal osmotic regulation in trpv4-/- mice, Proc. Natl. Acad. Sci. USA 100 (23), 13698–703, 2003. Mizuno, A., Matsumoto, N., Imai, M., and Suzuki, M., Impaired osmotic sensation in mice lacking TRPV4, Am. J. Physiol. Cell Physiol. 285 (1), C96–101, 2003. Caterina, M.J., Schumacher, M.A., Tominaga, M., Rosen, T.A., Levine, J.D., and Julius, D., The capsaicin receptor: a heat-activated ion channel in the pain pathway, Nature 389 (6653), 816–24, 1997. Jordt, S.E. and Julius, D., Molecular basis for species-speciﬁc sensitivity to ‘‘hot’’ chili peppers, Cell 108 (3), 421–30, 2002. Caterina, M.J., Rosen, T.A., Tominaga, M., Brake, A.J., and Julius, D., A capsaicinreceptor homologue with a high threshold for noxious heat, Nature 398 (6726), 436–41, 1999. Peier, A.M., Reeve, A.J., Andersson, D.A., Moqrich, A., Earley, T.J., Hergarden, A.C., Story, G.M., Colley, S., Hogenesch, J.B., McIntyre, P., Bevan, S., and Patapoutian, A., A heat-sensitive TRP channel expressed in keratinocytes, Science 296 (5575), 2046–49, 2002. Smith, G.D., Gunthorpe, M.J., Kelsell, R.E., Hayes, P.D., Reilly, P., Facer, P., Wright, J.E., Jerman, J.C., Walhin, J.P., Ooi, L., Egerton, J., Charles, K.J., Smart, D., Randall, A.D., Anand, P., and Davis, J.B., TRPV3 is a temperature-sensitive vanilloid receptor–like protein, Nature 418 (6894), 186–90, 2002. Xu, H., Ramsey, I.S., Kotecha, S.A., Moran, M.M., Chong, J.A., Lawson, D., Ge, P., Lilly, J., Silos-Santiago, I., Xie, Y., DiStefano, P.S., Curtis, R., and Clapham, D.E., TRPV3 is a calcium-permeable temperature-sensitive cation channel, Nature 418 (6894), 181–86, 2002. Voets, T., Droogmans, G., Wissenbach, U., Janssens, A., Flockerzi, V., and Nilius, B., The principle of temperature-dependent gating in cold- and heat-sensitive TRP channels, Nature 430 (7001), 748–54, 2004. Tominaga, M. and Caterina, M.J., Thermosensation and pain, J. Neurobiol. 61 (1), 3–12, 2004.

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50. Chung, M.K., Lee, H., and Caterina, M.J., Warm temperatures activate TRPV4 in mouse 308 keratinocytes, J. Biol. Chem. 278 (34), 32037–46, 2003. 51. Todaka, H., Taniguchi, J., Satoh, J., Mizuno, A., and Suzuki, M., Warm temperaturesensitive transient receptor potential vanilloid 4 (TRPV4) plays an essential role in thermal hyperalgesia, J. Biol. Chem. 279 (34), 35133–38, 2004. 52. Anishkin, A. and Kung, C., Microbial mechanosensation, Curr. Opin. Neurobiol. 15 (4), 397–405, 2005. 53. Kim, D., Physiology and pharmacology of two-pore domain potassium channels, Curr. Pharm. Des. 11 (21), 2717–36, 2005. 54. Earley, S., Heppner, T.J., Nelson, M.T., and Brayden, J.E., TRPV4 forms a novel Ca2+ signaling complex with ryanodine receptors and BKCa channels, Circ. Res. 97 (12), 1270–79, 2005. 55. Voets, T., Prenen, J., Vriens, J., Watanabe, H., Janssens, A., Wissenbach, U., Bodding, M., Droogmans, G., and Nilius, B., Molecular determinants of permeation through the cation channel TRPV4, J. Biol. Chem. 277 (37), 33704–10, 2002. 56. Kung, C. and Blount, P., Channels in microbes: so many holes to ﬁll, Mol. Microbiol. 53 (2), 373–80, 2004. 57. Liu, J. and Siegelbaum, S.A., Change of pore helix conformational state upon opening of cyclic nucleotide-gated channels, Neuron 28 (3), 899–909, 2000. 58. Zhen, X.G., Xie, C., Fitzmaurice, A., Schoonover, C.E., Orenstein, E.T., and Yang, J., Functional architecture of the inner pore of a voltage-gated Ca2+ channel, J. Gen. Physiol. 126 (3), 193–204, 2005. 59. Yeh, B.I., Kim, Y.K., Jabbar, W., and Huang, C.L., Conformational changes of pore helix coupled to gating of TRPV5 by protons, Embo J. 24 (18), 3224–34, 2005.

9

TRPV4: A Multifunctional Nonselective Cation Channel with Complex Regulation Tim D. Plant Philipps University

Rainer Strotmann Universität Leipzig

CONTENTS Introduction............................................................................................................125 Tissue Expression of TRPV4 ................................................................................126 Regulation of TRPV4 Activity ..............................................................................127 Regulation of TRPV4 by the Extracellular Osmolarity ...........................127 Mechanosensitivity of TRPV4 ..................................................................129 Mechanism of Activation by Hypotonic Solutions and Mechanical Stimuli.............................................................................130 Activation of TRPV4 by Phorbol Ester Derivatives.................................131 Regulation of TRPV4 by Temperature .....................................................132 Ca2+ Dependence of TRPV4 .....................................................................133 Interaction of Different Stimuli.................................................................135 Properties of Channels Formed by TRPV4 ..........................................................135 Biophysical Properties ...............................................................................135 Blockers .....................................................................................................136 Conclusions............................................................................................................136 References..............................................................................................................136

INTRODUCTION Mammalian TRP channels form a large family with around thirty members. From sequence similarity, TRPs can be divided into three major TRP subfamilies: the classical or canonical subfamily (TRPC), the melastatin-related subfamily (TRPM), 125

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and the vanilloid-receptor–related subfamily (TRPV).1 In addition, there are a number of more distantly related subfamilies: TRPA (ankyrin), TRPP (polycystin), and TRPML (mucolipidin).1–3 TRPV1, the ﬁrst member of the TRPV family and the sensory neuron receptor for vanilloid ligands like capsaicin, which is also responsive to noxious heat (>42°C), was found by expression cloning,4 as were the more distantly related epithelial Ca2+ channels TRPV55 and TRPV6.6 The other members, TRPV2,7 TRPV4 (see below), and later TRPV3,8–10 were found by homology screens. TRPV4 was found by screening expressed sequence tag databases for sequences with similarity to TRPV1, TRPV2, and the C. elegans TRPV isoform OSM-9. Lacking a consensus on nomenclature at the time, TRPV4 was given a variety of names—OTRPC4 (OSM-9-like TRP channel 4),11 VROAC (vanilloid receptor– related osmotically activated channel),12 TRP1213 and VRL-2 (vanilloid receptor– like channel 2)14—by the different groups who cloned the channel. TRPV4 has 871 amino acids, and structural features of the channel are intracellular N- and C-termini, six membrane spanning segments (S1–S6), a reentrant pore-forming loop between S5 and S6, and at least three ankyrin domains in the cytosolic N-terminus (see, e.g., Figure 9.2). Even though TRPV4 shows sequence similarity to other members of the TRPV family, particularly to TRPV1–3, a coexpression study has indicated that TRPV4 preferentially forms homomers,15 and, as yet, there is no evidence for heteromultimeric combinations with other TRPVs.

TISSUE EXPRESSION OF TRPV4 TRPV4 is widely expressed. TRPV4 is strongly expressed in the kidneys, and orthologues from most species (mouse, human, and rat) have been cloned from this tissue.11–14 In addition, the human orthologue has been cloned from a hypothalamus library and the chicken variant from an auditory epithelium library.12 In Northern hybridization, TRPV4 mRNA was detected in the heart, endothelium, brain, liver, placenta, lung, trachea, and salivary glands.11–14 mRNA is also present in airway smooth muscle16 and in the substantia nigra pars compacta.17 In the brain, in situ hybridization shows obvious mRNA expression in neurons of the circumventricular organs and in ependymal cells of the choroid plexus of the lateral and fourth, but not third, ventricles, and in scattered neurons in other regions of the brain.12 In the trigeminal ganglia and dorsal root ganglia, TRPV4 mRNA is present in large sensory neurons.12,14,18 In the inner ear, TRPV4 mRNA is present in inner and outer hair cells of the organ of Corti, and in hair cells of the semicircular canals and utriculae.12 TRPV4 mRNA is also present in the auditory ganglion and in marginal cells of the stria vascularis.12 In the respiratory tract, TRPV4 protein is expressed in the epithelia of the trachea and lung, submucosal glands, and mononuclear cells.14 Furthermore, protein was also detected in sympathetic ganglia and in sympathetic and parasympathetic nerve ﬁbers in a number of tissues.14 TRPV4 is also expressed in the hypothalamus and in keratinocytes,19 and in the epithelium of the oviduct.20 In the kidney tubules, TRPV4 expression is localized to constitutively or conditionally water impermeable (antidiuretic hormone-sensitive) segments,14,21 where it is mainly localized to the basolateral membrane.21

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REGULATION OF TRPV4 ACTIVITY Much of what is known about TRPV4 regulation comes from studies of the channel heterologously expressed in mammalian cells, although there are an increasing number of reports on a channel with similar properties in native tissues known to express TRPV4. Following heterologous expression, cells expressing TRPV4 often display spontaneous channel activity, which results in an elevated basal intracellular Ca2+ concentration ([Ca2+]i; Fig. 9.1A) and in spontaneous currents characteristic for TRPV4 (Fig. 9.1B,C). As yet unidentiﬁed intracellular factors are likely to be required for the maintenance of spontaneous channel activity as evidenced by the disappearance of currents within a few minutes in open whole cell patch-clamp (ruptured patch) recordings and the stable activity in perforated patch recordings. In the search for the mechanism of channel regulation, we tested a number of classical signaling pathways and cellular messengers but observed no clear increase or reduction in [Ca2+]i in cells expressing TRPV4 compared to control cells.11 Unlike TRPV1, TRPV4 is insensitive to capsaicin.11

REGULATION

OF

TRPV4

BY THE

EXTRACELLULAR OSMOLARITY

A number of cues indicated to investigators who ﬁrst studied TRPV4 that it may be an osmotically or mechanically sensitive channel. These included the knowledge that OSM-9 from C. elegans is involved in behavioral responses to increases in extracellular osmolarity and mechanical stimulation in the worm.22 Furthermore, some of the tissues where mammalian TRPV4 expression is found are exposed to changes in osmolarity (the kidney) or are involved in osmoregulation (the circumventricular organs) or mechanosensation (hair cells in the ear). Changes in osmolarity provide both an osmotic and mechanical stimulus, owing to cell swelling and membrane stretching. TRPV4 is highly sensitive to changes in extracellular osmolarity around the normal osmolarity of body ﬂuids. Reductions in the extracellular osmolarity result in increases in [Ca2+]i and membrane currents (Fig. 9.1A,B), whereas osmolarities above 300 mosmol l-1 decrease both [Ca2+]i and currents (Fig. 9.1C) in those cells displaying spontaneous activity.11 Signiﬁcant changes in Ca2+ inﬂux were observed in response to changes in extracellular osmolarity of as little as 1 percent.12 Stronger reductions in osmolarity result in larger increases in [Ca2+]i and currents, which are often not graded but show a slow increase followed by a very rapid response to a high level.11 This behavior probably reﬂects the positive feedback effect of Ca2+ entry on the channel (see below). In addition to relatively stable increases in [Ca2+]i, Liedtke et al. also reported that some cells respond to decreases in osmolarity with Ca2+ oscillations.12 Responses to reduced osmolarity show differences depending on the expression system, recording conditions, and laboratory. Gao et al.23 reported no elevation of basal Ca2+ in TRPV4-expressing cells and found very poor Ca2+ responses to hypotonic solutions at room temperature. Responses were nearly fourfold larger at 37°C. Others have observed strong [Ca2+]i responses to reduced osmolarity at room temperature.11,24 A more intact intracellular environment is probably important for the

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FIGURE 9.1 Responses of TRPV4-expressing HEK293 cells to the extracellular osmolarity and to the ligand 4αPMA. (A) [Ca2+]i measurements in TRPV4-expressing cells reveal an elevated basal [Ca2+]i in many cells indicative of spontaneous channel activity in those cells. Application of a hypotonic solution (200 mosmol l−1) resulted in large increases in [Ca2+]i in most cells that were reversible on returning to 300 mosmol l−1. Recording of the ﬂuorescence ratio F340/F380 from cells loaded with the Ca2+ indicator fura-2AM. (B) Currents recorded using the perforated patch technique from a HEK293 cell heterologously expressing TRPV4. Spontaneous currents were observed at 300 mosmol l−1, and reducing the osmolarity to 200 mosmol l−1 resulted in a transient increase in current followed by a decrease (“inactivation”) to levels lower than those before stimulation. (C) Spontaneous currents in TRPV4-expressing cells are decreased by raising the extracellular osmolarity from 300 to 320 mosmol l−1 in a perforated patch recording. (D) Activation of TRPV4 by the 4α-phorbol ester derivate 4αphorbol myristate acetate (4αPMA). Application of 4αPMA (1 μM) resulted in a rapid, transient increase in current mediated by TRPV4 in a ruptured patch whole-cell recording. Note the characteristic shape of the IV relation and the similarity in shape of the IV relation for spontaneous, osmotically activated, and 4αPMA-activated currents. The reversal potential in B differs from that in C and D owing to the reduced Na+ concentration in the hypotonic solution.

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response to hypotonicity. Current responses to hypotonic solutions are slow and weak in ruptured patch whole-cell recordings compared to those in perforated-patch recordings. In ruptured patch-clamp recordings, current responses to hypotonic solutions were also much smaller (ninefold) than responses to 4α-phorbol esters, potent ligands that activate TRPV4.25 Other groups report no response of TRPV4 to reduced osmolarity in CHO cells, but responses in HEK cells.26 Other factors that may inﬂuence the response to changes in osmolarity include proteins like microtubuleassociated protein 7 (MAP7) that interact with TRPV4.27 Consistent with the data on the osmosensitivity of heterologously expressed TRPV4, TRPV4-/- mice displayed defects in osmoregulation.28,29 TRPV4 has also been suggested to be involved in the nociceptive response of primary sensory neurons,18,30 and the [Ca2+]i response of airway smooth muscle cells to hypotonic stimulation.16 In endothelial cells, part of the response to hypotonic solutions is abolished in TRPV4-/- mice.31 In C. elegans lacking the TRPV isoform OSM-9, TRPV4 targeted to the appropriate sensory neurons restored responses to osmotic stimuli.32 Surprisingly, in this context, TRPV4 rescues responses to hypertonic solutions, whereas in mammalian cells, channel activation is seen on application of hypotonic solutions. The reason for this difference is unclear. Perhaps worm neurons, in contrast to mammalian cells, can generate an appropriate messenger to activate TRPV4 in response to hypertonic stimuli.

MECHANOSENSITIVITY

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TRPV4

The involvement of TRPV4 in responses to cell swelling and its localization in tissues like the inner ear, sensory neurons, and endothelial cells that are known to express mechanosensitive channels raise the question of whether TRPV4 is a mechanosensitive channel. We saw no activation of TRPV4 in response to membrane stretch by applying suction to the patch pipette in the cell-attached mode in HEK293 cells.11 A more recent study has, however, shown that TRPV4expressing cells respond in the whole-cell mode to cell inﬂation with a current activation.26 TRPV4’s responsiveness to noxious mechanical stimuli has also been suggested from a study of TRPV4-/- mice.26,28 TRPV4 also rescues mechanosensitivity in sensory neurons from C. elegans lacking OSM-9.32 TRPV4 was a major candidate for a mechanosensitive channel in hair cells in the cochlea involved in hearing. TRPV4−/− mice displayed a delayed onset hearing loss and an increased susceptibility to acoustic injury.33 The precise role of TRPV4 in hearing has not been identiﬁed, but the channel does not appear to be the mechanosensitive channel in hair cells. The expression of TRPV4 in tissues that show ﬂow- or shear stress-dependent Ca2+ entry like ﬂow-sensitive nephron segments and the vascular endothelium make TRPV4 a candidate for a channel sensitive to these stimuli. Increases in [Ca2+]i sensitive to ruthenium red, an inhibitor of TRPV channels (see below), have been reported in response to shear stress in HEK293 cells expressing TRPV4 at 37°C, but not at room temperature.23 To date, there has been no demonstration of TRPV4 involvement in responses to these stimuli in native tissues. An involvement in ﬂowsensitive Ca2+ increases in the nephron is not consistent with a report of a mainly

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basolateral localization in tubule cells.21 Surprisingly, increases in viscosity produced by adding dextran to the extracellular solution have been reported to increase currents in Hela cells expressing TRPV4.20

MECHANISM OF ACTIVATION AND MECHANICAL STIMULI

BY

HYPOTONIC SOLUTIONS

In the original demonstrations that TRPV4 could be regulated by changes in the extracellular osmolarity, it was unclear how the channel responds to these changes. We were careful to point out that the channel may respond directly to changes in osmolarity or may be activated by an endogenous signaling cascade sensitive to swelling.11 A delay of at least twenty seconds before TRPV4-expressing cells respond to osmotic stimuli, however, suggests that a slower process (e.g., cell swelling or signaling pathways activated by cell swelling) is involved in the response. Because many cellular signaling cascades have been demonstrated to be activated by changes in cell volume, there is no lack of prospective candidates.34 In contrast to volumeregulated anion channels, which are activated by a mechanism involving a swellinginduced reduction in intracellular ionic strength, and tyrosine kinase activation via small GTP-binding proteins, TRPV4 did not respond to infusion of GTPγS or to reductions in intracellular ionic strength.35 Nonreceptor tyrosine kinases have, however, been implicated in the response of TRPV4 to hypotonic solutions, and hypotonic stress has been reported to increase tyrosine phosphorylation of native and heterologously expressed TRPV4 by members of the Src family of tyrosine kinases.36 When coexpressed with TRPV4, Lyn, a tyrosine kinase of this family, dramatically increased phosphorylation of Tyr253 (Figure 9.2) in response to treatment with hypotonic

FIGURE 9.2 Transmembrane topology of TRPV4 showing the sites shown to be involved in the regulation of the channel and in determining channel properties. Abbreviations: SFK: src family kinase, Ank: Ankryrin domain, RR: ruthenium red, CaMBD: calmodulin-binding domain.

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solutions.36 Mutation of this tyrosine resulted in a loss of the Ca2+ response of TRPV4expressing cells to hypotonic solutions, leading to the suggestion that the response of TRPV4 to hypotonic cell swelling is mediated by tyrosine kinase-dependent phosphorylation. However, neither Vriens et al.24 nor we (unpublished data) could conﬁrm the loss of responsiveness to hypotonic solutions with this mutant. Watanabe et al. showed that TRPV4 can be activated by products of arachidonic acid breakdown.37 Arachidonic acid and endogenous activators of TRPV4, like the cannabinoid anadamide, that yield arachidonic acid are metabolized in a cytochrome P450 epoxygenase-dependent manner to epoxyeicosatrienoic acids (EETs). Inhibition of the cytochrome P450 epoxygenase strongly inhibited channel activation by arachidonic acid, and application of EETs activated heterologously expressed and endothelial TRPV4.31,37 Responses to hypotonic solutions were reduced by structurally unrelated inhibitors of PLA2 and by inhibitors of CYP450 epoxygenase24,31 and were increased by upregulation of the CYP450 epoxygenase.31 From these data, it is probable that the sensitivity to changes in extracellular osmolarity that TRPV4 confers on cells results from activation of an endogenous signaling cascade activated by changes in cell volume: swelling-induced activation of PLA2, release of arachidonic acid from membrane phospholipids, followed by CYP450 epoxygenasedependent metabolism of arachidonic acid to EETs, which then activate the channel. There is also evidence that, like the effects of hypotonic solutions, the activation of TRPV4 by increased extracellular viscosity is also PLA2 dependent.20 However, it remains unclear whether EETs activate TRPV4 by a direct interaction with the channel or whether other mediators are involved. A further interesting open question is how basal channel activity is generated and how reductions in [Ca2+]i and currents occur in response to hypertonic solutions. A simple explanation would be that PLA2 is active under isotonic conditions and that activity can be decreased by hypertonic solutions. However, Vriens et al. found that inhibitors of PLA2 have no effect on the basal activity of TRPV4.24 Thus, different mechanisms are likely to be involved in the generation of basal TRPV4 activity and in the response of the channel to hypotonic solutions.

ACTIVATION

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TRPV4

BY

PHORBOL ESTER DERIVATIVES

A very important ﬁnding for TRPV4 was the identiﬁcation of phorbol ester derivatives as channel activators (Figure 9.1D).25 TRPV4 was found to be activated by 4α-phorbol ester derivatives with EC50 values around 0.2 μM for 4α-phorbol didecanoate (4αPDD). These 4α-phorbol ester derivatives are commonly used as negative controls for the 4β-phorbol esters. The latter are widely used as activators of protein kinase C, (PKC) but also affect other proteins with C1-domains independently of PKC. The 4α-phorbol ester derivatives are not known to activate other ion channels and therefore represent speciﬁc tools to activate TRPV4 and study TRPV4-mediated effects. Under the conditions used in the study of Watanabe et al., the 4β-phorbol ester phorbol myristate acetate (PMA) was a less potent activator of TRPV4 than the 4α derivate, and acted as a partial agonist.25 In ruptured patch whole-cell experiments, 4α-phorbol esters are more effective than hypotonic solutions in activating TRPV4,25 but in more intact cells, [Ca2+]i increases in response to hypotonic solutions

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and 4αPDD are similar.24 The effects of 4α-phorbol ester derivatives are not mediated by the PLA2 pathway that is involved in activating TRPV4 by hypotonic solutions, because 4α-phorbol ester-mediated activation still occurs in the presence of PLA2 inhibitors.24 However, although the effects of hypotonic solutions and phorbol esters are mediated by different pathways, the effects of heat (see below) and 4αPDD seem to share a common mechanism. By analogy to TRPV1, for which a YS motif in the S2–S3 linker was shown to be involved in the binding of capsaicin,38 Vriens et al. identiﬁed a YS motif at the N-terminal end of S3 (Tyr555 and Ser556, Figure 9.2).24 Mutations of Tyr555 led to a loss of responsiveness to 4αPDD, but not to arachidonic acid, lending further support to the hypothesis that different pathways are involved in the responses to cell swelling and phorbol esters. Furthermore, the Tyr555 mutation also led to a loss of the response to heat (see below). The independence of pathways for the response of TRPV4 to cell swelling and arachidonic acid metabolism, and the responses to 4α-phorbol esters and heat have recently been conﬁrmed in endothelial cells.31 Studies performed at 37°C rather than at room temperature report different effects of 4β-phorbol ester derivatives.23,39 At the higher temperatures, 4β-phorbol ester derivatives had larger effects than at room temperature, and a large part of these effects was prevented by PKC inhibitors. These results suggest that the 4β-phorbol ester derivatives may have PKC-dependent and PKC-independent effects on TRPV4.

REGULATION

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TRPV4

BY

TEMPERATURE

Members of the TRPV family, with the exception of TRPV5 and TRPV6, form temperature-sensitive channels with temperature sensitivities ranging from warm (TRPV3/TRPV4) through hot (TRPV1), to very hot (TRPV2).40 TRPV4 has a temperature activation threshold between 25°C and 33°C in HEK293 cells,19,41 and around 27°C in Xenopus oocytes.19 Responses to warming show desensitization, but this is incomplete, leaving a spontaneously active current component at temperatures around 37°C. TRPV4 responses to heat are also observed in cell-attached patches, but not in cell-free patches,41 suggesting that cellular components not present or functional in excised patches are involved in the production of a messenger responsible for the heat response. Heat-activated currents are smaller than those in response to 4αPDD. The current density in response to warming to 38°C was about one ﬁfth of that in response to 4αPDD. However, there seem to be similarities in the responses to heat and 4αPDD because both responses are lost in the mutants of the YS motif in S3.24 This suggests that heat may inﬂuence the production of an as-yet-unidentiﬁed endogenous messenger that binds to a region of TRPV4 involving S3. Responses of TRPV4 to warm temperatures and its expression in sensory neurons, keratinocytes, and in the hypothalamus indicate a role for TRPV4 in thermosensation and thermoregulation. TRPV4-/- mice show decreased frequencies of nerve discharge in response to thermal stimulation, a decrease in the number of responsive ﬁbers, and longer latencies to escape from thermal stimuli in a model of thermal hyperalgesia.42 A lack of TRPV4 has no effect on escape latency in

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response to heat in the absence of hyperalgesia,26,28,42 with the exception of a study that described an increase in tail withdrawal latency to moderately hot temperatures.43 The latter study also showed that TRPV4-/- mice showed a preference for warmer ﬂoor temperatures. TRPV4 expression in the hypothalamus is an indicator of a possible role of TRPV4 in thermoregulation. However, Liedtke and Friedman found no difference in body temperature or differences in the temperature response to cold stress in TRPV4-/- mice.28

Ca2+ DEPENDENCE

OF

TRPV4

Most Ca2+-permeable channels show some feedback regulation by Ca2+ to prevent deleteriously large increases in [Ca2+]i or to shape the time course of channel activity. There is clear experimental evidence that, in addition to allowing Ca2+ to permeate, TRPV4 is closely regulated by [Ca2+]i. As described above, responses of TRPV4 to hypotonic solutions, phorbol esters, or heat are transient with rapid activation followed by inactivation/decay (Figures 9.1B, D). Ca2+ is involved both in the decaying phase and the activation phase of TRPV4; in the latter, it is not only a charge carrier, but also a potentiator of channel activity. Outward currents mediated by TRPV4 are strongly reduced by removing extracellular Ca2+. Similarly, removing extracellular Ca2+ abolishes spontaneous TRPV4 currents in parallel with the reduction in [Ca2+]i.44 There is a very close correlation between [Ca2+]i and the amplitude of TRPV4 currents. Replacement of extracellular Ca2+ by Ba2+ or Sr2+, which also permeate well (see below), leads to a reduction in spontaneous inward and outward currents, a result that indicates that these ions are unable to substitute for Ca2+ in the mechanism involved in generating spontaneous channel activity. In the absence of extracellular Ca2+, or when Ca2+ is replaced by Ba2+, activation of TRPV4 by hypotonic solutions or 4αPMA is slow, taking several minutes rather than seconds to reach a maximum. Ca2+ could either regulate the activity of TRPV4 by inﬂuencing Ca2+-dependent steps in signaling pathways leading to channel activation, or by binding to the channel or to channel-associated proteins. TRPV4 does not have an obvious Ca2+binding site like an EF hand, suggesting that Ca2+ does not bind directly to the channel. Many other TRP channels have calmodulin (CaM)-binding sites in their C-termini, but regulation via Ca2+ and CaM has only been convincingly demonstrated for a few isoforms, including the TRPV isoforms TRPV145 and TRPV6.46,47 We identiﬁed a CaM-binding site similar to that in the C-terminus of human TRPV646 in the intracellular C-terminus of TRPV4 (Figure 9.2).44 This binding site is a highly conserved helical stretch of amino acids, VGRLRRDRWSSVVPRV, starting at position 814. In in vitro CaM-binding experiments, mutations of amino acids within this region, including the point mutation W822A, prevented Ca2+-dependent CaM binding.44 For the C-terminal CaM-binding site of TRPV6, similar mutations that prevent Ca2+-dependent CaM binding decrease the rate of a slower phase of channel inactivation.46 In strong contrast to TRPV6, we found that the mutations that prevent CaM binding to C-terminal peptides of TRPV4 inﬂuence Ca2+-dependent potentiation and not inactivation of the channel.

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Wild-type TRPV4 shows strong potentiation on the addition of Ca2+ following channel activation by hypotonic solutions or 4α-phorbol esters in a Ca2+-free medium.44 After transient expression, the full-length channels with mutations that prevent Ca2+ -CaM binding showed no potentiation of current responses to 4αPMA on Ca2+ addition, and only slow activation by 4αPMA in the presence of extracellular Ca2+. The speed of activation in Ca2+ resembled that of TRPV4-wt in the absence of Ca2+ or when Ba2+ replaced Ca2+. From these data, we suggest that there is a Ca2+-dependent component of TRPV4 activation. Irrespective of the primary activator, hypotonic medium, lipophilic ligands, or heat, Ca2+ entering through TRPV4 binds to CaM, and Ca2+-CaM interacts with the C-terminal binding domain, resulting in positive feedback activation of TRPV4, increasing both the speed and amplitude of the response. It remains unclear, however, how Ca2+-CaM binding exerts its effects on channel activity and what the physiological role of this mechanism may be. If unchecked, positive feedback involving Ca2+ could lead to large, potentially harmful increases in [Ca2+]i and, thus, some kind of inhibitory mechanism is necessary to turn the channel off. Ca2+ limits the entry of Ca2+ through other Ca2+permeable channels by feedback inhibition. Likewise, Ca2+ is involved in the decay of TRPV4 currents and in controlling channel availability. Large, fast TRPV4 current responses decay rapidly to levels below those of spontaneous currents before activation (see, e.g., Figure 9.1B), even in the continuous presence of the activating stimulus. Current responses in the absence of Ca2+ are in general smaller, and display slower activation kinetics (as described above) but also much weaker current decay, even when current amplitudes similar to those in the presence of Ca2+ are attained. With increasing extracellular [Ca2+], current responses to 4α-phorbol esters activate more rapidly (as expected from the Ca2+-dependent potentiation described above), become smaller in amplitude, and decay more completely and rapidly.48 Watanabe et al.48 also saw a reduction in current amplitude as [Ca2+]i was raised via the intracellular solution in the patch pipette, with an IC50 of around 600 nM. The mechanism by which Ca2+ inactivates TRPV4 remains unclear. Other TRPV channels (TRPV1 and TRPV6) have been shown to inactivate in a Ca2+-dependent fashion. TRPV6 shows biphasic inactivation kinetics, with a fast component in the millisecond and a slow component in the second range. The fast component was shown to depend on a sequence motif in the ﬁrst intracellular loop,49 while the slow component is mediated by Ca2+-CaM binding to a C-terminal binding domain.46 Both sites have no homologues in the TRPV4 sequence. By analogy to TRPV1, however, for which a tyrosine residue in S6 has been shown to be involved in Ca2+ permeation and Ca2+-dependent desensitization,50 Watanabe et al. found that mutation of a phenylalanine residue at position 707 in S6 of TRPV4 (Figure 9.2) to alanine resulted in a reduction and slowing of current decay but did not modify the sensitivity to [Ca2+]i.48 In the same study, mutation of Glu797 in the C-terminal tail (Figure 9.2) increased spontaneous channel activity, but the increase in [Ca2+]i in response to 4αPDD was similar.48 Glu797 is close to the CaM-binding domain, and it remains to be seen whether the mutation of this residue inﬂuences CaM binding. Thus, Ca2+-dependent inactivation of TRPV4 seems to rely on structural features that have not yet been identiﬁed.

TRPV4: A Multifunctional Nonselective Cation Channel

INTERACTION

OF

135

DIFFERENT STIMULI

Because a number of mechanisms converge on TRPV4, they are likely to interact and inﬂuence one another. Liedtke et al. found that responses of TRPV4 to decreases in osmolarity were larger at 37°C than at room temperature.12 As described above for the individual activators, Gao et al. found that responses to most activators, including 4αPDD, PMA, hypotonic solutions, and shear stress, are increased at 37°C compared to room temperature.23 For the response of TRPV4 to temperature, raising the osmolarity very signiﬁcantly depressed responses to heat in HEK293 cells and oocytes, whereas decreasing the osmolarity initiated a response alone and potentiated the response to heat in HEK293 cells.19 Somewhat surprisingly, however, the effects of 5,6-EETs, unlike the effects of hypotonic solutions,12,23 were found not to be affected by heat and were reduced in hypoosmotic conditions.51

PROPERTIES OF CHANNELS FORMED BY TRPV4 BIOPHYSICAL PROPERTIES TRPV1 to TRPV4 differ not only in their sequences but also in their biophysical properties from TRPV5 and TRPV6. Many of the differences in properties result from differences in the Ca2+ permeability of the isoforms. The IV relation of TRPV4 has a characteristic shape, showing outward rectiﬁcation in Ca2+-containing solutions, but also an increase in conductance, and concomitantly in inward current, at negative membrane potentials (Figure 9.1B, C, D). The reversal potential in normal extracellular solutions is just positive to 0 mV (Figure 9.1C, D). The outward rectiﬁcation results from a block by extracellular Ca2+. Lowering the extracellular [Ca2+] increases inward currents and leads to a loss of rectiﬁcation in Ca2+-free solutions.52 Asp672 and Asp682 in the pore region (Figure 9.2) are involved in the inhibition of TRPV4 currents by extracellular Ca2+. Neutralization of both leads to a strong reduction in inhibition and a linearization of the IV relation.52 Interestingly, the IV shape is similar for single-channel and whole-cell currents. Because of the outward rectiﬁcation, the single-channel conductance for outward currents (88–100 pS) is larger than that for inward currents (30–60 pS). A value of 310 pS at +80 mV, a value much larger than that in other studies, was obtained in a cursory investigation of single-channel properties in another study.12 In studies other than the latter, the relatively large variability in the conductance values obtained for inward currents probably results from differences in the solutions bathing the external face of the patch. Lower values were obtained in more physiological Ca2+ concentrations11 than in Ca2+-free solutions,37,41 as expected from the effect of Ca2+ on whole-cell currents.52 Thus, where studied carefully, data on single-channel conductance are consistent, and variabilities in the conductance for inward currents result from the use of different experimental conditions. The reversal potential of around +5 to +10 mV for TRPV4 with standard extraand intracellular solutions and abolition of most of the inward current on replacement of extracellular cations with the large cation N-methyl-D-glucamine (NMDG+) indicates that the channel is a nonselective cation channel. TRPV4 is moderately Ca2+ permeable with PCa/PNa values of between 6 and 10.11,25 Other divalent cations also

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permeate with a PMg/PNa of 2–3, a PSr/PNa of 9, and a PBa/PNa of 0.7–7.25,44,52 The channel discriminates poorly among monovalent cations with PK:PCs:PNa:PLi = 2:1.3:1:0.9.25,35 Mutations that affect Ca2+ inhibition and the shape of the IV relation also moderately reduce the Ca2+ permeability.52 Mutation of Met680, which, by analogy to K+ channels, is located at the center of the selectivity ﬁlter (Figure 9.2), leads to a reduction in current and a strong reduction in Ca2+ permeability.52

BLOCKERS Studies on TRP channels suffer from the lack of speciﬁc pharmacological modulators. Like other members of the TRPV subfamily, TRPV4 is blocked by extracellular ruthenium red (RR) in micromolar concentrations.11,25,52 The block by RR is voltage dependent, with inward currents being more strongly inhibited than outward currents. It is also notable that inward currents are inhibited much more rapidly than outward currents. For native currents mediated by TRPV4 in keratinocytes, but not in HEK293 cells, an increase in outward current on RR application has been reported.53 In our hands, application of 10 μM RR to TRPV4 in HEK293 cells resulted in an increase in outward current followed by inhibition (unpublished observations). Neutralization of Asp682 (Figure 9.2), a residue involved in the block of TRPV4 by extracellular Ca2+ (see above), shifted the potential dependence of and slowed the block by RR.52 Thus, Asp682 is likely to be in the outer region of the pore accessible to the large cation. TRPV4 is also inhibited by micromolar Gd3+ 11 and SKF96365,32 which, however, inhibit many other TRP channels and native cation channels in a similar concentration range and cannot be considered speciﬁc.

CONCLUSIONS A number of the TRPV channels are likely to have highly speciﬁc roles, and their expression is restricted to a very limited number of tissues. TRPV4 shows a wider expression pattern, with expression in cells with a variety of physiological functions from sensory transduction to epithelial transport. In addition, TRPV4 shares with TRPV1 the property of being regulated by a surprising variety of stimuli. For TRPV4, these include physical stimuli like osmotic stress, mechanical stress, and heat, as well as endogenous lipid agonists. Unlike many channels that serve the same speciﬁc functional role in different tissues, data emerging for TRPV4 suggest that the physiological stimulus may depend on the tissue context and that a combination of stimuli will determine the activity of the channel.

REFERENCES 1. Clapham, D.E., Montell, C., Schultz, G., and Julius, D., International Union of Pharmacology. XLIII. Compendium of voltage-gated ion channels: transient receptor potential channels, Pharmacol. Rev. 55, 591–596, 2003. 2. Moran, M.M., Xu, H., and Clapham, D.E., TRP ion channels in the nervous system, Curr. Opin. Neurobiol. 14, 362–369, 2004.

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3. Pedersen, S.F., Owsianik, G., and Nilius, B., TRP channels: An overview, Cell Calcium 38, 233–252, 2005. 4. Caterina, M., Schumacher, M.A., Tominaga, M., Rosen, T.A., Levine, J.D., and Julius, D., The capsaicin receptor: a heat-activated ion channel in the pain pathway., Nature 389, 816–824, 1997. 5. Hoenderop, J.G.J., van der Kemp, A.W.C.M., Hartog, A., van de Graaf, S.F.J., van Os, C.H., Willems, P.H.G.M., and Bindels, R.J.M., Molecular identiﬁcation of the apical Ca2+ channel in 1,25-dihydroxyvitamin D3-responsive epithelia, J. Biol. Chem. 274, 8375–8378, 1999. 6. Peng, J.B., Chen, X.Z., Berger, U.V., Vassilev, P.M., Tsukaguchi, H., Brown, E.M., and Hediger, M.A., Molecular cloning and characterization of a channel-like transporter mediating intestinal calcium absorption, J. Biol. Chem. 274, 22739–22746, 1999. 7. Caterina, M.J., Rosen, T.A., Tominaga, M., Brake, A.J., and Julius, D., A capsaicin receptor homologue with a high threshold for noxious heat, Nature 398, 436–441, 1999. 8. Smith, G.D., Gunthorpe, M.J., Kelsell, R.E., Hayes, P.D., Reilly, P., Facer, P., Wright, J.E., Jerman, J.C., Walhin, J.P., Ooi, L., Egerton, J., Charles, K.J., Smart, D., Randall, A.D., Anand, P., and Davis, J.B., TRPV3 is a temperature-sensitive vanilloid receptorlike protein, Nature 418, 186–190, 2002. 9. Xu, H., Ramsey, I.S., Kotecha, S.A., Moran, M.M., Chong, J.A., Lawson, D., Ge, P., Lilly, J., Silos-Santiago, I., Xie, Y., DiStefano, P.S., Curtis, R., and Clapham, D.E., TRPV3 is a calcium-permeable temperature-sensitive cation channel, Nature 418, 181–186, 2002. 10. Peier, A.M., Reeve, A.J., Andersson, D.A., Moqrich, A., Earley, T.J., Hergarden, A.C., Story, G.M., Colley, S., Hogenesch, J.B., McIntyre, P., Bevan, S., and Patapoutian, A., A heat-sensitive TRP channel expressed in keratinocytes, Science 296, 2046–2049, 2002. 11. Strotmann, R., Harteneck, C., Nunnenmacher, K., Schultz, G., and Plant, T.D., OTRPC4, a nonselective cation channel that confers sensitivity to extracellular osmolarity, Nat. Cell Biol. 2, 695–702, 2000. 12. Liedtke, W., Choe, Y., Marti-Renom, M.A., Bell, A.M., Denis, C.S., Sali, A., Hudspeth, A.J., Friedman, J.M., and Heller, S., Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor, Cell 103, 525–535, 2000. 13. Wissenbach, U., Bödding, M., Freichel, M., and Flockerzi, V., Trp12, a novel Trp-related protein from kidney, FEBS Lett. 485, 127–134, 2000. 14. Delany, N.S., Hurle, M., Facer, P., Alnadaf, T., Plumpton, C., Kinghorn, I., See, C.G., Costigan, M., Anand, P., Woolf, C.J., Crowther, D., Sanseau, P., and Tate, S.N., Identiﬁcation and characterization of a novel human vanilloid receptor-like protein, VRL-2, Physiol. Genomics 4, 165–174, 2001. 15. Hellwig, N., Albrecht, N., Harteneck, C., Schultz, G., and Schaefer, M., Homo- and heteromeric assembly of TRPV channel subunits, J. Cell Sci. 118, 917–928, 2005. 16. Jia, Y., Wang, X., Varty, L., Rizzo, C.A., Yang, R., Correll, C.C., Phelps, P.T., Egan, R.W., and Hey, J.A., Functional TRPV4 channels are expressed in human airway smooth muscle cells, Am. J. Physiol. Lung Cell Mol. Physiol. 287, L272–278, 2004. 17. Guatteo, E., Chung, K.K., Bowala, T.K., Bernardi, G., Mercuri, N.B., and Lipski, J., Temperature sensitivity of dopaminergic neurons of the substantia nigra pars compacta: involvement of TRP channels, J. Neurophysiol. 94, 3069–3080, 2005.

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18. Alessandri-Haber, N., Yeh, J.J., Boyd, A.E., Parada, C.A., Chen, X., Reichling, D.B., and Levine, J.D., Hypotonicity induces TRPV4-mediated nociception in rat, Neuron 39, 497–511, 2003. 19. Güler, A.D., Lee, H., Iida, T., Shimizu, I., Tominaga, M., and Caterina, M., Heatevoked activation of the ion channel, TRPV4, J. Neurosci. 22, 6408–6414, 2002. 20. Andrade, Y.N., Fernandes, J., Vazquez, E., Fernandez-Fernandez, J.M., Arniges, M., Sanchez, T.M., Villalon, M., and Valverde, M.A., TRPV4 channel is involved in the coupling of ﬂuid viscosity changes to epithelial ciliary activity, J. Cell Biol. 168, 869–874, 2005. 21. Tian, W., Salanova, M., Xu, H., Lindsley, J.N., Oyama, T.T., Anderson, S., Bachmann, S., and Cohen, D.M., Renal expression of osmotically responsive cation channel TRPV4 is restricted to water-impermeant nephron segments, Am. J. Physiol. Renal Physiol. 287, F17–24, 2004. 22. Colbert, H.A., Smith, T.L., and Bargmann, C.I., Osm-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation and olfactory adaptation in Caenorhabditis elegans, J. Neurosci. 17, 8259–8269, 1997. 23. Gao, X., Wu, L., and O’Neil, R.G., Temperature-modulated diversity of TRPV4 channel gating: activation by physical stresses and phorbol ester derivatives through protein kinase C-dependent and -independent pathways, J. Biol. Chem. 278, 27129–27137, 2003. 24. Vriens, J., Watanabe, H., Janssens, A., Droogmans, G., Voets, T., and Nilius, B., Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4, Proc. Natl. Acad. Sci. USA 101, 396–401, 2004. 25. Watanabe, H., Davis, J.B., Smart, D., Jerman, J.C., Smith, G.D., Hayes, P., Vriens, J., Cairns, W., Wissenbach, U., Prenen, J., Flockerzi, V., Droogmans, G., Benham, C.D., and Nilius, B., Activation of TRPV4 channels (hVRL-2/mTRP12) by phorbol derivatives, J. Biol. Chem. 277, 13569–13577, 2002. 26. Suzuki, M., Mizuno, A., Kodaira, K., and Imai, M., Impaired pressure sensation in mice lacking TRPV4, J. Biol. Chem. 278, 22664–22668, 2003. 27. Suzuki, M., Hirao, A., and Mizuno, A., Microtubule-associated protein 7 increases the membrane expression of transient receptor potential vanilloid 4 (TRPV4), J. Biol. Chem. 278, 51448–51453, 2003. 28. Liedtke, W. and Friedman, J.M., Abnormal osmotic regulation in trpv4-/- mice, Proc. Natl. Acad. Sci. USA 100, 13698–13703, 2003. 29. Mizuno, A., Matsumoto, N., Imai, M., and Suzuki, M., Impaired osmotic sensation in mice lacking TRPV4, Am. J. Physiol. Cell Physiol. 285, C96–101, 2003. 30. Alessandri-Haber, N., Dina, O.A., Yeh, J.J., Parada, C.A., Reichling, D.B., and Levine, J.D., Transient receptor potential vanilloid 4 is essential in chemotherapyinduced neuropathic pain in the rat, J. Neurosci. 24, 4444–4452, 2004. 31. Vriens, J., Owsianik, G., Fisslthaler, B., Suzuki, M., Janssens, A., Voets, T., Morisseau, C., Hammock, B.D., Fleming, I., Busse, R., and Nilius, B., Modulation of the Ca2+ permeable cation channel TRPV4 by cytochrome P450 epoxygenases in vascular endothelium, Circ. Res. 97, 908–915, 2005. 32. Liedtke, W., Tobin, D.M., Bargmann, C.I., and Friedman, J.M., Mammalian TRPV4 (VR-OAC) directs behavioral responses to osmotic and mechanical stimuli in Caenorhabditis elegans, Proc. Natl. Acad. Sci. USA 100 Suppl. 2, 14531–14536, 2003. 33. Tabuchi, K., Suzuki, M., Mizuno, A., and Hara, A., Hearing impairment in TRPV4 knockout mice, Neurosci. Lett. 382, 304–308, 2005.

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34. Lang, F., Busch, G.L., Ritter, M., Volkl, H., Waldegger, S., Gulbins, E., and Haussinger, D., Functional signiﬁcance of cell volume regulatory mechanisms, Physiol. Rev. 78, 247–306, 1998. 35. Nilius, B., Prenen, J., Wissenbach, U., Bödding, M., and Droogmans, G., Differential activation of the volume-sensitive cation channel TRP12 (OTRPC4) and volumeregulated anion currents in HEK-293 cells, Pﬂügers Arch. 443, 227–233, 2001. 36. Xu, H., Zhao, H., Tian, W., Yoshida, K., Roullet, J.B., and Cohen, D.M., Regulation of a transient receptor potential (TRP) channel by tyrosine phosphorylation. SRC family kinase-dependent tyrosine phosphorylation of TRPV4 on TYR-253 mediates its response to hypotonic stress, J. Biol. Chem. 278, 11520–11527, 2003. 37. Watanabe, H., Vriens, J., Prenen, J., Droogmans, G., Voets, T., and Nilius, B., Anandamide and arachidonic acid use epoxyeicosatrienoic acids to activate TRPV4 channels, Nature 424, 434–438, 2003. 38. Jordt, S.E. and Julius, D., Molecular basis for species-speciﬁc sensitivity to ‘‘hot’’ chili peppers, Cell 108, 421–430, 2002. 39. Xu, F., Satoh, E., and Iijima, T., Protein kinase C-mediated Ca2+ entry in HEK 293 cells transiently expressing human TRPV4, Br. J. Pharmacol. 140, 413–421, 2003. 40. Patapoutian, A., Peier, A.M., Story, G.M., and Viswanath, V., ThermoTRP channels and beyond: mechanisms of temperature sensation, Nat. Rev. Neurosci. 4, 529–539, 2003. 41. Watanabe, H., Vriens, J., Suh, S.H., Benham, C.D., Droogmans, G., and Nilius, B., Heat-evoked activation of TRPV4 channels in an HEK293 cell expression system and in native mouse aorta endothelial cells, J. Biol. Chem. 277, 47044–47051, 2002. 42. Todaka, H., Taniguchi, J., Satoh, J., Mizuno, A., and Suzuki, M., Warm temperaturesensitive transient receptor potential vanilloid 4 (TRPV4) plays an essential role in thermal hyperalgesia, J. Biol. Chem. 279, 35133–35138, 2004. 43. Lee, H., Iida, T., Mizuno, A., Suzuki, M., and Caterina, M.J., Altered thermal selection behavior in mice lacking transient receptor potential vanilloid 4, J. Neurosci. 25, 1304–1310, 2005. 44. Strotmann, R., Schultz, G., and Plant, T.D., Ca2+-dependent potentiation of the nonselective cation channel TRPV4 is mediated by a C-terminal calmodulin-binding site, J. Biol. Chem. 278, 26541–26549, 2003. 45. Numazaki, M., Tominaga, T., Takeuchi, K., Murayama, N., Toyooka, H., and Tominaga, M., Structural determinant of TRPV1 desensitization interacts with calmodulin, Proc. Natl. Acad. Sci. USA 100, 8002–8006, 2003. 46. Niemeyer, B.A., Bergs, C., Wissenbach, U., Flockerzi, V., and Trost, C., Competitive regulation of CaT-like-mediated Ca2+ entry by protein kinase C and calmodulin, Proc. Natl. Acad. Sci. USA 98, 3600–3605, 2001. 47. Lambers, T.T., Weidema, A.F., Nilius, B., Hoenderop, J.G., and Bindels, R.J., Regulation of the mouse epithelial Ca2+ channel TRPV6 by the Ca2+-sensor calmodulin, J. Biol. Chem. 279 (28), 28855–28861, 2004. 48. Watanabe, H., Vriens, J., Janssens, A., Wondergem, R., Droogmans, G., and Nilius, B., Modulation of TRPV4 gating by intra- and extracellular Ca2+, Cell Calcium 33, 489–495, 2003. 49. Nilius, B., Prenen, J., Hoenderop, J.G., Vennekens, R., Hoefs, S., Weidema, A.F., Droogmans, G., and Bindels, R.J., Fast and slow inactivation kinetics of the Ca2+ channels ECaC1 and ECaC2 (TRPV5 and 6): role of the intracellular loop located between transmembrane segment 2 and 3, J. Biol. Chem. 277, 30852–30858, 2002.

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50. Mohapatra, D.P., Wang, S.Y., Wang, G.K., and Nau, C., A tyrosine residue in TM6 of the vanilloid receptor TRPV1 involved in desensitization and calcium permeability of capsaicin-activated currents, Mol. Cell Neurosci. 23, 314–324, 2003. 51. Nilius, B., Vriens, J., Prenen, J., Droogmans, G., and Voets, T., TRPV4 calcium entry channel: a paradigm for gating diversity, Am. J. Physiol. Cell Physiol. 286, C195–205, 2004. 52. Voets, T., Prenen, J., Vriens, J., Watanabe, H., Janssens, A., Wissenbach, U., Bödding, M., Droogmans, G., and Nilius, B., Molecular determinants of permeation through the cation channel TRPV4, J. Biol. Chem. 277, 33704–33710, 2002. 53. Chung, M.K., Lee, H., Mizuno, A., Suzuki, M., and Caterina, M.J., TRPV3 and TRPV4 mediate warmth-evoked currents in primary mouse keratinocytes, J. Biol. Chem. 279, 21569–21575, 2004.

10

TRPV4 and TRPM3 as Volume-Regulated Cation Channels Christian Harteneck and Günter Schultz Institut für Pharmakologie

CONTENTS Introduction............................................................................................................141 TRPV4 ...................................................................................................................142 Molecular Features and Tissue Expression...............................................142 Activation and Pharmacological Properties ..............................................143 Regulation and Cellular Function .............................................................144 TRPM3...................................................................................................................145 Molecular Features and Tissue Expression...............................................145 Activation and Pharmacological Properties ..............................................146 Summary ................................................................................................................147 Acknowledgments..................................................................................................148 References..............................................................................................................148

INTRODUCTION Changes of the extracellular osmolarity result in swelling or shrinkage of cells by increases or decreases in intracellular water along the ionic concentration gradient, thereby changing cell volume. For the viability of the cells, fast adaptation to the extracellular tonicity is essential and, therefore, a variety of cellular mechanisms protects cells from osmotic stress. By modulation of ion channels and transporters, cells modulate their intracellular ion concentrations in order to adapt to environmental conditions and facilitate cellular functions.1,2 The change in cell volume is accompanied by reorganization of the cytoskeleton, involving the structure-forming proteins as well as proteins involved in the regulation of cell architecture. Despite the variety of systems described to be involved in volume regulation like potassium and chloride channels, osmolytes, and others, the connections between the different systems are as unclear as the proteins sensing changes in tonicity and subsequent signaling cascades.

141

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We recently characterized two different TRP-homologous cation channels, TRPV4 and TRPM3, as proteins mediating calcium entry in cells upon extracellular application of hypotonic solutions.3,4 Proteins of the TRP family as integral membrane proteins form pores in the lipid bilayer of the plasma membrane, regulating ion ﬂuxes across the membrane. Today the TRP superfamily is a superfamily of proteins subdivided into at least seven different subfamilies. High sequence similarity in the region of the pore-forming domains is the common feature of the classic (TRPC), melastatin-like (TRPM), and vanilloid-like (TRPV) subfamilies’ functional properties, and proposed topology is the common theme of all TRP channels.5–7 The cDNAs of two orphan proteins, named TRPV4 and TRPM3 according to sequence similarity and order of appearance, were cloned by their sequence similarity to Drosophila TRP. The proteins transiently expressed in HEK293 cells could be characterized as nonselective cation channels mediating calcium entry upon extracellular application of hypoosmolar solutions. This chapter summarizes the known data for TRPV4 and TRPM3.

TRPV4 MOLECULAR FEATURES

AND

TISSUE EXPRESSION

The mRNA coding for TRPV4 is transcribed from the gene locus chromosome 12q24.1 in humans, 12q16 in rats, and 5 in mice. The gene is organized by twelve exons separated by intronic sequences. The 3.25 kb mRNA encodes an open reading frame resulting in a protein of 876 amino acids for the regularly expressed form.4,8–10 In addition to this form, a shorter variant was identiﬁed by RT-PCR approaches. The proposed open reading frame would result in a protein of 811 amino acids.11 Like other TRP-homologous channels, the predicted topology argues for transmembrane proteins with six membrane-spanning domains and intracellular N- and C-terminal ends (Figure 10.1). The ion-permeable pore is formed by the ﬁfth- and sixth-membrane spanning domains and the sequence between, forming a loop and contributing to the pore in analogy to the structure of the bacterial

FIGURE 10.1 Transmembrane topology and functional domains of TRPV4.

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potassium channel Ksc. The sequence of the cytoplasmic N-terminus encodes three ankyrin repeat domains, a feature found in the sequences of all TRP channels of the TRPC and TRPV subfamilies. The ankyrin repeat domains in TRP channels are discussed to function as a protein–protein-interaction domain, linking members of the TRPC and TRPV family to the cytoskeleton or allowing multimerization of channel complexes.12 The role of the three ankyrin repeat domains in TRPV4 is still unclear. However, it has been shown that TRPV4 is tyrosine phosphorylated at tyrosine in position 253 located in the ﬁrst ankyrin repeat domain, allowing speculations that protein–protein interactions of TRPV4 via the ﬁrst ankyrin repeat domain may be dynamically regulated by tyrosine phosphorylation.13 The functional data and the controversy about Y253 phosphorylation and functional consequences are summarized in the next chapter.13,14 On the other hand, in the sequence of the TRPV4 short splice variant, the third ankyrin repeat domain is Cterminally deleted, allowing speculations about whether targeting complex formation or function of TRPV4 is modulated by splice processes.11 The highest expression level of TRPV4 is found in the kidney.4 The renal expression of TRPV4 is restricted to the epithelial cells of the water-impermeable part of the nephron, the thin and thick ascending limbs (ATL, TAL), the distal convolute tubule (DCT), and the connecting tubule.15 Lower TRPV4 expression levels were found in many other tissues like the skin, trachea, liver, lung, blood vessels, and the brain. On the cellular level, TRPV4 expression has been reported for keratinocytes, airway smooth muscle cells, aortic myocytes, many epithelial cell types (lining the alveoli, submucosal glands, and salivary glands), endothelial cells, neurons of the circumventricular organs (VOLT, SFO, MnPO, and choroids plexus), neurosensory cells, inner-ear hair cells, sensory neurons, Merkel cells, and sympathetic and parasympathetic nerve ﬁbers.8–11,15–18 Because TRPV4 is expressed in such a great variety of cell types, it likely functions as a polymodal sensory protein with speciﬁed functions, depending on the cell type, expressing TRPV4 together with different associated proteins. Data from knockout mice revealed that TRPV4 is involved in sensation of osmotic, mechanic, and heat stress (see below).

ACTIVATION

AND

PHARMACOLOGICAL PROPERTIES

TRPV4 activity is modulated by extracellular osmolarity. In heterologous expression systems, high plasma membrane levels of TRPV4 are obtained, resulting in an increased basal calcium concentration due to the large amount of active channels according to the law of mass action. TRPV4 is permeable for calcium and sodium with a permeability ratio PCa/PNa 6 and a conductance of 90 pS.19 Activity is blocked by ruthenium red (10 μM) and gadolinium or lanthanum ions (100 μM).19 The reduction of the basal calcium concentration by the application of hyperosmolar solutions (330 mosmol/l) shows that TRPV4 is inhibited by the hyperosmolarity, whereas hypoosmolar solutions (200–250 mosmol/l) activate TRPV4.4,8 Applying hypotonic solutions to HEK293 cells was associated with cell swelling. However, the activation of TRPV4 was independent of pressure applied. Later it had been shown that TRPV4 activation is independent of temperature. In a temperature range of 24°C to 40°C, heat activates and sensitizes TRPV4 to shear stress.20,21 As a

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polymodal sensor protein, TRPV4 mediates calcium entry by ligands like 4α− stereomeric forms of phorbol esters and by arachidonic acid and anandamide.22,23 Experiments using inhibitors of the cytochrome P450 epoxygenase activity demonstrate that TRPV4 activation is not directly mediated by arachidonic acid and anandamide. TRPV4 is rather activated by the arachidonic acid metabolite epoxyeicosatrienoic acid.23,24

REGULATION

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CELLULAR FUNCTION

TRPV4 is involved in volume regulation. Attempts to clarify the physiologic role of TRPV4 by genetic inactivation of TRPV4 in mice show impairments of volume balance in two independently generated mouse lines.25,26 Conﬂicts arise from conﬂicting descriptions of drinking behavior, serum osmolarity, and the basal serum level of the antidiuretic hormone. In both reports, hyperosmotic challenge of the TRPV4-/- mice induced an increase in the serum concentration of the antidiuretic hormone. TRPV4 is not only involved in volume balance but also in nociceptive transduction.27–29 TRPV-/- mice show reduced sensitivity to pressure of the tail and acidic nociception.29 In hot-plate tests, the latency to escape following hyperalgesia induced by carrageenan was longer in TRPV4-/- mice.28 Additionally, TRPV4-/- mice show an altered thermal selection behavior.27 Wild-type mice prefer 30°C, whereas TRPV4 knockout mice choose 34°C.27 In hearing tests, elderly TRPV4-/- mice show hearing impairment with higher thresholds for auditory brainstem responses, whereas the hearing loss after acoustic overexposure is delayed in TRPV4-/- mice. This ﬁnding corresponds with the ﬁnding that TRPV4 maps to 12q21–24, a locus associated with dominant nonsyndromic hereditary hearing impairment in humans.30 Parallel to ﬁnding the physiological role of TRPV4, the molecular characteristics of this channel protein have been clariﬁed. The activity of TRPV4 is negatively regulated by increased intracellular calcium concentrations. The calcium sensor protein calmodulin binds to the calmodulin-binding site of TRPV4 in a calciumdependent manner, decreasing TRPV4 activity by an auto-regulatory mechanism (see Figure 10.1).31 The calcium/calmodulin-binding site is located C-terminally to the putative sixth membrane-spanning domain. While the calmodulin-binding site negatively regulates TRPV4, the phosphorylation of tyrosine Y253 and the microtubule-associated protein 7 (MAP-7) have been described as positive regulators of TRPV4 activity.13,14,32 MAP-7 binds to TRPV4 in the region between the putative sixth membrane and the calcium/calmodulin-binding domain, linking TRPV4 to the cytoskeletal network and thereby increasing membrane localization and current density.32 Whereas the MAP-7 interaction modulates targeting of TRPV4 to the plasma membrane, retrieval back from the surface to the intracellular compartment of the Golgi network and the endoplasmic reticulum is assumed to be mediated by PACS proteins.33 By co-immunoprecipitation, it has been shown that TRPV4 interacts with PACS-1 proteins. PACS proteins recognize acidic clusters in channel proteins and mediate retrieval of the proteins from the plasma membrane (see Figure 10.1). Therefore, it is likely that PACS mediate retrieval of TRPV4 in a phosphorylationdependent manner as it has been shown for the polycystin channel protein PKD2 (TRPP2). While the kinase phosphorylating an amino acid in the acidic cluster of

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TRPV4 (174 to 188) is unknown, TRPV4 is phosphorylated by src kinases.13 Kinases of the src family phosphorylate TRPV4 at position Y253 in a hypotonicity-dependent manner, providing an activation mechanism for TRPV4. However, whether or not tyrosine phosphorylation of TRPV4 is necessary for the hypotonicity-induced activation of TRPV4 is controversially discussed. While David Cohen and coworkers showed loss of function by changing tyrosine 253 to alanine, Bernd Nilius et al. found normal hypotonic stimulation independent of the type of amino acid at position 253.13,14 Previous reports describing accumulation and release of arachidonic acid upon hypotonic stimulation make it likely that calcium entry upon hypotonic induction is mediated in an arachidonic acid–dependent manner via TRPV4 activation by epoxyeicosatrienoic acid.23,24,34

TRPM3 MOLECULAR FEATURES

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TISSUE EXPRESSION

The mRNAs coding for TRPM3 are transcribed from the gene locus chromosome 9q21.11–12 in humans, 1q51 in rats, and 19 in mice. The open frame of TRPM3 is organized in at least 30 exons separated by one of the longest intronic sequences in chromosome 9.35 Available information of transcribed mRNAs reveals that many different mRNA species coding for a variety of TRPM3 proteins are transcribed. We initially cloned a TRPM3 variant coded by 1,325 amino acids from mice and later from humans. We expressed the corresponding cDNA in HEK293 cells and were able to functionally characterize TRPM as a nonselective cation channel.3,36 In Northern blot analyses of mice brains, we found at least three transcripts of different lengths, whereas Lee et al. cloned six variants from human kidneys, which vary in short deletions and insertions indistinguishable by Northern blot analysis.3,37 Figure 10.2 summarizes the differences in the TRPM3 proteins found so far by

FIGURE 10.2 Transmembrane topology and sequence modiﬁcations by splice events of TRPM3.

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analyses of the sequence databases. The variability of TRPM3 transcripts depends on the presence of two or probably three alternative start positions and two different C-terminal ends. The differences in C-terminal ends depend on a probably regulated splicing effect of a highly conserved splice site found in the long variants at amino acid position 1318. In the mRNAs coding for the high molecular weight variants, the splicing machinery ignored this site, whereas the mRNA species coding for the low molecular weight variants resulted from recognition of this site and the addition of an accessory exon coding for eight amino acids and carrying the 3´-untranslated region by the splice machinery. At this time, it is unclear whether mRNA species of all possible variations or a set of three to ﬁve mRNAs with deﬁned changes in the primary sequence are transcribed in a cell type–speciﬁc manner. Further analysis of the translated proteins by Western blot analyses and other biochemical methods will answer these questions. In our initial analyses, we detected up to four distinct protein forms in Western blot analyses under denaturing conditions. Starting from the sequence information of an EST from a kidney, we cloned the complete cDNAs coding for TRPM3 from a human fetal brain and human kidney. Both tissues represent the major expression site of TRPM3.3 In the kidney, TRPM3 is localized to the collecting tubular epithelium in the medulla, medulla rays, and periglomerular regions. Low levels were found in other epithelia like the proximal convoluted tubular epithelium.37 By RT-PCR, we detected lower TRPM3 levels in the brain, ovary, and pancreas. In the brain, TRPM3 is expressed in the cerebellum, plexus choroideus, locus coerulus, posterior hypothalamus, and substantia nigra.3,37,38

ACTIVATION

AND

PHARMACOLOGICAL PROPERTIES

Summarizing the four reports on functional expression of TRPM3, one can conclude that the differences in lengths of TRPM3 mRNAs and corresponding proteins probably result in different activation mechanisms.3,36–38 While Lee and coworkers as well as Oberwinkler et al. studied long variants encoded by 1,545 and 1,725 amino acids, respectively, we functionally characterized a shorter variant of 1,325 amino acids. The short variant forms a channel protein of approximately 150 kDa in humans and mice, whereas the long variant corresponds to proteins of 180 kDa in Western blot analyses. Heterologously expressed in HEK293 cells, TRPM3 (the 1325aa-variant) forms a calcium-permeable pore in the plasma membrane mediating calcium entry.3 In untreated cells, the constitutive activity of expressed TRPM3 results in increased intracellular calcium concentrations when compared to control-transfected cells. The TRPM3 pore is also permeable for manganese, allowing manganese-quenching experiments that show that the activity can be blocked by lanthanum and gadolinium ions and by 2-APB, whereas the channel blocker SK&F-96365 is ineffective.39 Like in experiments of heterologously expressed TRPV4, the spontanous activity was blocked by applying hypertonic solutions, whereas enhanced intracellular calcium concentrations were measured upon extracellular application of hypotonic solutions. The expression of TRPM3 in the kidney and the activation by hypotonicity argue for the function of TRPM3 in renal osmo-homeostasis. Recently, we characterized TRPM3 as the ﬁrst cation channel activated by sphingosine.36 In transiently transfected HEK293 cells, sphingosine and its precursor

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dihydrosphingosine induce TRPM3 currents. Other metabolites of the sphingolipid pathway like ceramides and sphingosine-1-phosphate, described as a ligand of G-protein-coupled receptors, were ineffective in activating TRPM3. The responses to sphingosine are restricted to TRPM3, while sphingosine was ineffective in HEK293 cells expressing TRPC3, TRPC4, TRPC5, TRPV4, TRPV5, TRPV6, or TRPM2. The sphingosine-induced, TRPM3-mediated calcium entry is not affected by store depletion of calcium or by compounds inhibiting protein kinase C. Therefore, a direct activation of TRPM3 by sphingosine is very likely. The intracellular sphingosine concentration depends on de novo synthesis, degradation of sphingolipids by ceramidases, activity of sphingosine kinases and sphingosine-1-phosphatases, and degradation of sphingosine by sphingosine lyases. It is unclear which pathway results in sphingosine concentrations capable of inducing TRPM3 currents. Although we found no activation of TRPM3 by store-depletion protocols, Lee and coworkers characterized their TRPM3 variant to be activated by store-depletion protocols.37 They show results of calcium depletion and readdition protocols, but it is unclear whether the differences in activity of control- and TRPM3-transfected cells originate from activation of TRPM3 by store depletion or from spontaneous activity. Constitutive activity of long TRPM3 variants was also characterized by Oberwinkler et al. They described TRPM3 as a channel protein permeable for calcium and magnesium.38 They characterized the splice variation of TRPM3 already identiﬁed by Lee et al. with ten additional amino acids in the extracellular loop next to the putative sixth membrane-spanning segment.37,38 This insertion in TRPM3 corresponds to a stretch of seven amino acids in the published sequence of TRPM1. The presence of the additional amino acids results in a loss of ion permeability of the expressed TRPM3 protein and argues for a dominant-negative function for this variant or the function of this TRPM3 variant as a scaffolding protein.

SUMMARY The kidney as an organ controlling water and blood volume expresses both osmoregulated TRP-homologous channels, TRPM3 and TRPV4. Along the epithelial lining of the nephrone, TRPM3 and TRPV4 are found in different segments: TRPM3 in the proximal convoluted tubule and the collecting duct, and TRPV4 in the waterimpermeable segments of the thin and thick ascending limbs, the distal convolute tubule, and the connecting tubule. Based on the osmolarity-dependent activation, it is likely that both channel proteins are involved in the regulation of renal volume homeostasis, probably by establishing or modulating the osmotic gradient along the cortical-medullary axis. The complementary distribution in water-permeable and water-impermeable epithelials as well as the differential activation by sphingolipids and eicosanoids suggests that the channel proteins are integrated in different cellular signaling cascades necessary to maintain the speciﬁc architecture in the kidney. Besides the high expression in the kidney, both proteins are widely distributed in different tissues and cell types. As already summarized, TRPV4 is involved in a variety of functions (e.g., transduction of nociceptive, acoustic, and mechanic stimuli). In contrast to TRPV4, the characterization of TRPM3 is at the beginning.

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Currently, the data postulate that TRPM3 is involved in renal volume regulation and is integrated in sphingolipid signaling.

ACKNOWLEDGMENTS This work of the authors was supported by the Deutsche Forschungsgemeinschaft, Fonds der Chemischen Industrie and Sonnenfeld-Stiftung.

REFERENCES 1. Lang, F. et al., Functional signiﬁcance of cell volume regulatory mechanisms. Physiol. Rev. 78, 247–306, 1998. 2. Wehner, F., Olsen, H., Tinel, H., Kinne-Saffran, E., & Kinne, R.K., Cell volume regulation: osmolytes, osmolyte transport, and signal transduction. Rev. Physiol. Biochem. Pharmacol. 148, 1–80, 2003. 3. Grimm, C., Kraft, R., Sauerbruch, S., Schultz, G., & Harteneck, C., Molecular and functional characterization of the melastatin-related cation channel TRPM3. J. Biol. Chem. 278, 21493–501, 2003. 4. Strotmann, R., Harteneck, C., Nunnenmacher, K., Schultz, G., & Plant, T.D., OTRPC4, a nonselective cation channel that confers sensitivity to extracellular osmolarity. Nat. Cell Biol. 2, 695–702, 2000. 5. Harteneck, C., Plant, T.D., & Schultz, G., From worm to man: three subfamilies of TRP channels. Trends Neurosci. 23, 159–66, 2000. 6. Montell, C. et al., A uniﬁed nomenclature for the superfamily of TRP cation channels. Mol. Cell 9, 229–31, 2002. 7. Montell, C., The TRP superfamily of cation channels. Sci. STKE 2005, re3, 2005. 8. Liedtke, W. et al., Vanilloid receptor–related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor. Cell 103, 525–35, 2000. 9. Wissenbach, U., Bödding, M., Freichel, M., & Flockerzi, V., Trp12, a novel Trprelated protein from kidney. FEBS Lett. 485, 127–34, 2000. 10. Delany, N.S. et al., Identiﬁcation and characterization of a novel human vanilloid receptor–like protein, VRL-2. Physiol. Genomics 4, 165–74, 2001. 11. Arniges, M., Vazquez, E., Fernandez-Fernandez, J.M., & Valverde, M.A., Swellingactivated Ca2+ entry via TRPV4 channel is defective in cystic ﬁbrosis airway epithelia. J. Biol. Chem. 279, 54062–68, 2004. 12. Erler, I., Hirnet, D., Wissenbach, U., Flockerzi, V., & Niemeyer, B.A., Ca2+-selective transient receptor potential V channel architecture and function require a speciﬁc ankyrin repeat. J. Biol. Chem. 279, 34456–63, 2004. 13. Xu, H. et al., Regulation of a transient receptor potential (TRP) channel by tyrosine phosphorylation. SRC family kinase-dependent tyrosine phosphorylation of TRPV4 on TYR-253 mediates its response to hypotonic stress. J. Biol. Chem. 278, 11520–27, 2003. 14. Vriens, J. et al., Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4. Proc. Natl. Acad. Sci. USA 101, 396–401, 2004. 15. Tian, W. et al., Renal expression of osmotically responsive cation channel TRPV4 is restricted to water-impermeant nephron segments. Am. J. Physiol. Renal Physiol. 287, F17–24, 2004.

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16. Chung, M.K., Lee, H., & Caterina, M.J., Warm temperatures activate TRPV4 in mouse 308 keratinocytes. J. Biol. Chem. 278, 32037–46, 2003. 17. Guler, A.D. et al., Heat-evoked activation of the ion channel, TRPV4. J. Neurosci. 22, 6408–14, 2002. 18. Jia, Y. et al., Functional TRPV4 channels are expressed in human airway smooth muscle cells. Am. J. Physiol. Lung Cell Mol. Physiol. 287, L272–78, 2004. 19. Clapham, D.E., Montell, C., Schultz, G., & Julius, D., International Union of Pharmacology. XLIII. Compendium of voltage-gated ion channels: transient receptor potential channels. Pharmacol. Rev. 55, 591–96, 2003. 20. Gao, X., Wu, L., & O'Neil, R.G., Temperature-modulated diversity of TRPV4 channel gating: activation by physical stresses and phorbol ester derivatives through protein kinase C–dependent and –independent pathways. J. Biol. Chem. 278, 27129–37, 2003. 21. Watanabe, H. et al., Heat-evoked activation of TRPV4 channels in a HEK293 cell expression system and in native mouse aorta endothelial cells. J. Biol. Chem. 277, 47044–51, 2002. 22. Watanabe, H. et al., Activation of TRPV4 channels (hVRL-2/mTRP12) by phorbol derivatives. J. Biol. Chem. 277, 13569–77, 2002. 23. Watanabe, H. et al., Anadamide and arachidonic acid use epoxyeicosatrienoic acid to activate TRPV4 channels. Nature 424, 434–38, 2003. 24. Vriens, J. et al., Modulation of the Ca2+-permeable cation channel TRPV4 by cytochrome P450 epoxygenases in vascular endothelium. Circ. Res. 2005. 25. Mizuno, A., Matsumoto, N., Imai, M., & Suzuki, M., Impaired osmotic sensation in mice lacking TRPV4. Am. J. Physiol. Cell Physiol. 285, C96–101, 2003. 26. Liedtke, W. & Friedman, J.M., Abnormal osmotic regulation in trpv4-/- mice. Proc. Natl. Acad. Sci. USA 100, 13698–703, 2003. 27. Lee, H., Iida, T., Mizuno, A., Suzuki, M., & Caterina, M.J., Altered thermal selection behavior in mice lacking transient receptor potential vanilloid 4. J. Neurosci. 25, 1304–10, 2005. 28. Todaka, H., Taniguchi, J., Satoh, J., Mizuno, A., & Suzuki, M., Warm temperaturesensitive transient receptor potential vanilloid 4 (TRPV4) plays an essential role in thermal hyperalgesia. J. Biol. Chem. 279, 35133–38, 2004. 29. Suzuki, M., Mizuno, A., Kodaira, K., & Imai, M., Impaired pressure sensation in mice lacking TRPV4. J. Biol. Chem. 278, 22664–68, 2003. 30. Tabuchi, K., Suzuki, M., Mizuno, A., & Hara, A., Hearing impairment in TRPV4 knockout mice. Neurosci. Lett. 382, 304–8, 2005. 31. Strotmann, R., Schultz, G., & Plant, T.D., Ca2+-dependent potentiation of the nonselective cation channel TRPV4 is mediated by a C-terminal calmodulin binding site. J. Biol. Chem. 278, 26541–49, 2003. 32. Suzuki, M., Hirao, A., & Mizuno, A., Microtubule-associated protein 7 increases the membrane expression of transient receptor potential vanilloid 4 (TRPV4). J. Biol. Chem. 278, 51448–53, 2003. 33. Köttgen, M. et al., Trafﬁcking of TRPP2 by PACS proteins represents a novel mechanism of ion channel regulation. Embo J. 24, 705–16, 2005. 34. Basavappa, S., Pedersen, S.F., Jorgensen, N.K., Ellory, J.C., & Hoffmann, E.K., Swelling-induced arachidonic acid release via the 85-kDa cPLA2 in human neuroblastoma cells. J. Neurophysiol. 79, 1441–49, 1998. 35. Humphray, S.J. et al., DNA sequence and analysis of human chromosome 9. Nature 429, 369–74, 2004.

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36. Grimm, C., Kraft, R., Schultz, G., & Harteneck, C., Activation of the melastatin-related cation channel TRPM3 by D-erythro-sphingosine. Mol. Pharmacol. 67, 798–805, 2005. 37. Lee, N. et al., Expression and characterization of human transient receptor potential melastatin 3 (hTRPM3). J. Biol. Chem. 278, 20890–97, 2003. 38. Oberwinkler, J., Lis, A., Giehl, K.M., Flockerzi, V., & Philipp, S.E., Alternative splicing switches the divalent cation selectivity of TRPM3 channels. J. Biol. Chem. 280, 22540–48, 2005. 39. Xu, S.Z. et al., Block of TRPC5 channels by 2-aminoethoxydiphenyl borate: a differential, extracellular, and voltage-dependent effect. Br. J. Pharmacol. 145, 405–14, 2005.

11

TRPA1 : A Sensory Channel of Many Talents Marilia Z. P. Guimaraes Universidade Federal do Rio de Janeiro

Sven-Eric Jordt Yale University School of Medicine

CONTENTS Abstract ..................................................................................................................151 Introduction............................................................................................................152 TRPA1 in the Sensory Neural Response to Mustard Oil .....................................152 Mustard Oil and Capsaicin as Chemical Probes for the Pain Pathway .......................................................................................152 TRPA1 Mediates Mustard Oil Effects in Sensory Neurons .....................153 Activation of TRPA1 by Garlic Derivatives and Hazardous Unsaturated Aldehydes ..............................................................................154 Activation of TRPA1 by Cannabinoids.................................................................155 TRPA1 in Cold Sensation: Questions and Answers .............................................157 TRPA1 Is a Receptor-Operated Channel Acting in Concert with TRPV1 ...........................................................................................................158 TRPA1 as a Multiple Irritant Sensor.....................................................................159 Conclusion .............................................................................................................159 References..............................................................................................................160

ABSTRACT In mammals, TRPA1 is the sole member of the TRPA gene subfamily. Recent reports identiﬁed TRPA1 as a target for the noxious and inﬂammatory irritant mustard oil in peripheral sensory neurons, implicating a functional role in pain and neurogenic inﬂammation. Other studies suggest that TRPA1 participates in additional sensory processes, such as cold sensation and hearing. In this chapter, we summarize and discuss these recent ﬁndings and speculate about the potential physiological role of TRPA1 in chemosensation and pain transduction.

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INTRODUCTION TRPA1 is a member of the TRPA branch of the TRP ion channel gene family. On the structural level, TRPA channels are characterized by multiple N-terminal ankyrin repeats (~14 in the N-terminus of human TRPA1). Superﬁcially, TRPA channels resemble TRPN channels that were implicated in mechanotransduction and hearing in Drosophila and zebraﬁsh. However, the ion channel domain of TRPA channels is evolutionarily distant from TRPN channels. While other animals express two or more TRPA genes, mammals have only a sole TRPA gene, TRPA1. Far from having only a rudimentary presence, TRPA1 has attracted signiﬁcant attention from different areas of sensory research. In mammals, TRPA1 is expressed in a subset of peripheral sensory neurons, implicating a specialized role in sensory transduction. Recent studies found evidence that TRPA1 is involved in sensory neural responses to mustard oil, allicin, and other chemical irritants. Moreover, TRPA1 may serve as a sensor for noxious cold temperature. Other studies identiﬁed TRPA1 as a candidate for the auditory hair cell transduction channel. The potential role of TRPA1 in hearing will be discussed in a different chapter in this volume. Here, we focus on TRPA1 as a target for chemical sensory irritants and discuss its physiological role in acute and inﬂammatory pain.

TRPA1 IN THE SENSORY NEURAL RESPONSE TO MUSTARD OIL MUSTARD OIL AND CAPSAICIN AS CHEMICAL PROBES FOR THE PAIN PATHWAY Mustard and pungent roots such as wasabi or radish have been staples in the human diet for millennia. The pungent ingredient in these and other plants of the genus Brassica (the cabbages) is mustard oil (allyl isothiocyanate). Mustard oil is produced from a chemical precursor, sinigrin, by the enzyme myrosinase that is activated when mustard seeds are crushed in the presence of water. Besides inducing acute pain and irritation, mustard oil is also a potent inﬂammatory agent. Applying it to the skin induces reddening, swelling, edema, and plasma extravasation, accompanied by thermal and mechanical hyperalgesia, the painful hypersensitivity to otherwise innocuous thermal and mechanical stimuli. Since it was discovered that mustard oil activated inﬂammation through a neurogenic mechanism, mustard oil has became an invaluable tool in the study of the pain pathway.1 Mustard oil induces neurogenic inﬂammation by triggering the release of neuropeptides such as CGRP (calcitonin gene-related peptide) and Substance P from sensory nerve endings.2 These peptides activate local dilation and permeabilization of the vasculature and promote inﬁltration of the affected tissue by neutrophils and other immune cells.3 Mustard oil–induced neurogenic inﬂammation closely resembles neurogenic inﬂammation caused by capsaicin, the pungent ingredient in chili peppers.4 Capsaicin is a vanilloid compound and is structurally distinct from isothiocyanates. Both capsaicin and mustard oil were crucial for the discovery and characterization of a speciﬁc subset of nociceptive inﬂammatory sensory neurons, the C-ﬁbers. Pretreatment of the skin with

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capsaicin reversibly abolishes the induction of pain and inﬂammation by mustard oil, indicating that both compounds target the same populations of neurons. Psychophysical tests in human subjects also demonstrate a close relationship between the actions of capsaicin and mustard oil. When the human tongue is pretreated with capsaicin, placement of a mustard oil–soaked ﬁlter disk on the tongue failed to induce painful sensations.5 When newborn rats were injected with a large dose of capsaicin, they became insensitive to both capsaicin and mustard oil throughout their life spans.6 Injection of capsaicin causes permanent ablation of a subpopulation of sensory neurons with small soma diameters in all sensory ganglia.7,8 These neurons give rise to unmyelinated sensory ﬁbers, the C-ﬁbers, which are characterized by their sensitivity to capsaicin. Although mustard oil and capsaicin have similar effects, the pharmacology of capsaicin is much better deﬁned. The discovery of the high-afﬁnity capsaicin receptor ligand resiniferatoxin enabled the localization of capsaicin receptor sites on sensory neurons. Synthetic agonists and antagonists such as olvanil and capsazepine facilitated further detailed pharmacological and electrophysiological studies of vanilloid action. The cDNA encoding for the capsaicin receptor, TRPV1, was cloned in 1997, allowing receptor studies on the molecular level.9 Deleting the TRPV1 gene in mice revealed that TRPV1 is essential for inﬂammatory thermal hyperalgesia and that it contributes to heat- and acid-evoked pain.10,11 These and many other studies afﬁrmed the crucial role of TRPV1 in pain transduction, and TRPV1 has become one of the major drug discovery targets for the development of new analgesics. In contrast, very little was known about potential targets for mustard oil on sensory neurons. In most studies, mustard oil was topically applied or injected in pure form or as an oil emulsion. Speciﬁc antagonists or analogues with higher potencies were unknown, and for some time it was thought that mustard oil might either exert its effects through nonspeciﬁc chemical damage of sensory neurons or through activation of the capsaicin receptor. However, TRPV1-deﬁcient mice retained normal sensitivity to mustard oil,10 and TRPV1 is not activated by mustard oil in vitro.12 Indicating a more speciﬁc action, a few select studies showed that isothiocyanates induce neurogenic effects at much lower concentrations. In one study, mustard oil induced contraction of the rat bladder at micromolar concentrations through a neurogenic mechanism.13 Interestingly, this activity was blocked by ruthenium red, a cation channel blocker that also blocks TRP channels such as TRPV1. In a different study, benzyl isothiocyanate, a pungent structural congener of mustard oil, caused the neurogenic relaxation of precontracted arteries at micromolar concentrations.14 Similar to the discovery of the capsaicin receptor, the detection and analysis of the molecular targets for mustard oil in sensory neurons may devise new strategies for the development of analgesics and anti-inﬂammatory agents.

TRPA1 MEDIATES MUSTARD OIL EFFECTS

IN

SENSORY NEURONS

To investigate the cellular and molecular basis of mustard oil action, we studied responses of cultured rat sensory neurons to mustard oil by ratiometric calcium imaging. We found that mustard oil activated inﬂux of Ca2+ into ~35 percent of sensory neurons.12 All mustard oil–responsive neurons were also sensitive to capsaicin.

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Mustard oil–induced Ca2+ inﬂux could be blocked by ruthenium red. Thus, we hypothesized that mustard oil activates a ruthenium red–sensitive Ca2+-permeable ion channel in TRPV1-expressing neurons. Intriguingly, these characteristics were shared by the ion channel TRPA1, previously discovered as a potential mediator of responses to noxious cold stimuli in sensory neurons.15 TRPA1 is expressed in a subset of peptidergic TRPV1-positive neurons. Moreover, TRPA1 channel currents are sensitive to ruthenium red.15 These coincidences encouraged us to test responses of human and rat TRPA1 channels to mustard oil. We found that mustard oil and other pungent isothiocyanates induced robust ruthenium red–sensitive currents in Xenopus oocytes and in cultured mammalian cells expressing human and rat TRPA1. The potencies of mustard oil for TRPA1 activation and for sensory neural responses were comparable.12 Similarly, extracts from mustard seeds and wasabi activated TRPA1. Mouse TRPA1 was also found to be sensitive to mustard oil.16,17 Our recent analysis of mice with a targeted deletion in the TRPA1 gene indicates that TRPA1 may represent the sole site for the pungent and inﬂammatory action of mustard oil. TRPA1-deﬁcient mice do not display acute pain-related behavior after application of mustard oil to paws. Mustard oil–induced mechanical and thermal hyperalgesia were absent. Moreover, mustard oil failed to activate inﬂux of Ca2+ into TRPA1-deﬁcient cultured sensory neurons, indicating that TRPA1 may represent the sole site for the pungent and inﬂammatory action of these agents.

ACTIVATION OF TRPA1 BY GARLIC DERIVATIVES AND HAZARDOUS UNSATURATED ALDEHYDES In addition to mustard oil, TRPA1 is activated by other pungent plant products such as cinnamaldehyde, eugenol, gingerol, and methyl salicylate.16 However, very high concentrations of these compounds are needed to activate TRPA1. Two recent studies showed that extracts from garlic activate TRPA1.18,19 Garlic contains a variety of pungent organosulfur compounds, including allicin, a thiosulﬁnate compound, and diallyl disulﬁde. Thiosulﬁnates and pungent disulﬁdes are also present in onions and other plants of the genus allium. Thiosulﬁnates and allyl disulﬁde are structurally related to mustard oil, sharing allyl groups and labile carbon-sulfur bonds. Both allicin and allyl disulﬁde were found to be potent activators of TRPA1.18,19 Allicin and diallyl disulﬁde activate Ca2+ inﬂux into a subset of capsaicin-sensitive neurons that are also sensitive to mustard oil.18,19 Cultured neurons from TRPA1-deﬁcient mice failed to respond to allicin with Ca2+ uptake, conﬁrming that TRPA1 is the sole site of action of pungent organosulfur compounds. Garlic extracts and derivatives have hypotensive properties in vitro and in vivo in animal models. Adventitial sensory nerve ﬁbers in arteries contribute to vasodilation by activity-dependent release of the vasodilatory peptide CGRP. In mesenteric arterial preparations, garlic extracts induced vasodilation of precontracted arterial segments through a neurogenic pathway.18 Allicin, diallyl disulﬁde, and mustard oil had the same effects.18 Their activities were abolished by pretreatment of the preparation with capsaicin and by the TRP channel blocker ruthenium red. Capsaicin depletes CGRP from capsaicin-sensitive neurons, leaving subsequent neural activation

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ineffective. Taken together, these results suggest that allicin, diallyl disulﬁde, and mustard oil induce vasodilation in vitro by activating TRPA channels on capsaicinsensitive perivascular sensory nerve endings. Clearly, future pharmacological and genetic experiments are required to clarify whether this mechanism contributes to the systemic hypotensive effects of garlic in vivo. It remains controversial whether dietary garlic has beneﬁcial cardiovascular effects in humans. In addition to pungent organosulfur compounds, TRPA1 is activated by noxious unsaturated aldehydes.20 One of these aldehydes is acrolein (2-propenal), a potent lachrymator and pulmonary agent that was used in chemical warfare in the First World War. Acrolein is an environmental hazard produced during combustion, and it can be found in cigarette smoke, smoke from ﬁres, and smog. Moreover, acrolein is a toxic by-product of cyclophosphamide chemotherapy, causing hemorrhagic cystitis through neurogenic effects. While acrolein triggers Ca2+ inﬂux into cultured neurons from wild-type mice, neurons of TRPA1-deﬁcient mice fail to respond to this noxious compound.20

ACTIVATION OF TRPA1 BY CANNABINOIDS The Cannabis plant has been cultivated for centuries both for the production of hemp ﬁber and for its presumed medicinal and psychoactive properties. The bestknown effects of cannabinoids are changes in mood and motivation, as well as in appetite. However, cannabinoids also induce a complex mixture of dose-dependent cardiovascular effects that can cause postural hypertension or hypotension in humans.21 These effects revealed important roles for cannabinoids in the cardiovascular system. The major active ingredient in cannabis is Δ-9-tetrahydrocannabinol (THC), which produces most of the characteristic pharmacological effects. The identiﬁcation of this and other phytocannabinoids and their corresponding receptors suggested that endogenous substances, endocannabinoids, might exist that are capable of eliciting similar actions. Anandamide, a polyunsaturated fatty acid amide, was discovered as a potent endocannabinoid that activates cannabinoid receptors in the brain. It is thought that most cannabinoid actions happen through the activation of two cannabinoid receptors, CB1 and CB2, both G-protein-coupled receptors. CB1 was ﬁrst identiﬁed in the brain, where it is expressed abundantly. CB1 usually couples to Gi, leading to a reduction in cAMP and to Ca2+ channel inhibition and K+ channel stimulation. The second receptor, CB2, is expressed in immune system cells. Both receptors have been implicated in the cardiovascular actions of cannabinoids.22 In anesthetized rats, a bolus injection of anandamide induces a triphasic cardiovascular response, consisting of an initial transient fall in blood pressure and heart rate (phase I), followed by a pressor response (phase II), and culminating with prolonged hypotension (phase III).22 The third phase can be blocked by a synthetic cannabinoid antagonist, SR141716A (rimonabant), indicating that CB1 is involved. However, phase I could not be blocked by the same antagonist. In addition, in studies with isolated mesenteric arteries, it has been demonstrated that anandamide has powerful vasodilatory effects, whereas other synthetic cannabinoids known to activate CB1 do not. Furthermore, the vasodilator activity of anandamide in the perfused mesenteric vascular bed remains intact in CB1 knockout mice.23

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N

Mustard oil (Allyl isothiocyanate)

S

C

O S

S

S

S

Allicin (garlic)

Diallyl disulﬁde (garlic and onions)

O

O

Cinnamaldehyde

Acrolein

FIGURE 11.1 Diverse chemical activators of TRPA1: TRPA1 channels were initially identiﬁed as the target of mustard oil (allyl isothiocyanate), the pungent ingredient in mustard. Subsequently other pungent organosulfur compounds, such as allicin and diallyl disulﬁde, found in garlic and onions, were shown to activate TRPA1. Noxious aldehydes such as cinnamaldehyde and acrolein (2-propenal) also activate TRPA1.

The discovery that this effect was endothelium independent suggested that anandamide might act through a neurogenic mechanism. Indeed, anandamide was found to induce vasodilation by releasing CGRP from perivascular sensory neurons.23 Because this effect was blocked by ruthenium red and capsazepine, it was proposed that anandamide might stimulate TRPV1 receptors present in sensory nerves, which was shown to be the case.24 This report showed that endocannabinoids can act through receptors other than CB1 and CB2, and it identiﬁed a TRP channel as an ionotropic cannabinoid receptor. THC is known to cause hypotension in laboratory animals, even though in humans it might have the opposite effect.21 These differences might be due to the dose and/or experimental setting, such as anesthetized versus nonanesthetized subjects. In rat isolated mesenteric and hepatic arteries, THC induces extensive vessel relaxation.25 Similar to anandamide, THC was shown to act via a neurogenic, CB1and CB2-independent pathway. This effect is mediated by capsaicin-sensitive neurons, as depletion of CGRP by capsaicin abolished the vasodilatory actions of THC. Similar to the effect of anandamide, THC-induced vasodilation was sensitive to ruthenium red. However, THC was still able to induce vasodilation in arteries prepared from TRPV1 knockout mice, indicating that a target different from TRPV1 mediates this activity. This result led to the pursuit of yet another cannabinoid

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receptor with characteristics of a TRP channel. In subsequent experiments, it was found that THC activates Ca2+ inﬂux into a capsaicin- and mustard oil–sensitive population of sensory neurons. Because of TRPA1’s coexpression with TRPV1, its sensitivity to mustard oil, and its sensitivity to ruthenium red, it was tested as a potential receptor for THC. Indeed, THC activated cationic currents in oocytes and mammalian cells expressing TRPA1. This indicates that TRPA1 might be responsible for the vasodilatory actions of THC in isolated vessels and establishes TRPA1 as an additional ionotropic cannabinoid receptor. Although anandamide does not promote TRPA1 activity, these results suggest that other endogenous cannabinoid-like compounds exist that may modulate TRPA1 activity.

TRPA1 IN COLD SENSATION: QUESTIONS AND ANSWERS The physiological role of TRPA1 in sensory transduction is currently a matter of spirited debate. Initially, TRPA1 was identiﬁed as a potential mediator of noxious cold stimuli in nociceptive sensory neurons.15 This hypothesis was based on the observation that TRPA1 responds to cold stimuli in heterologous expression systems. However, other recent studies did not observe cold sensitivity of TRPA1 channels.12,17 Thus, it remains to be determined whether TRPA1 has intrinsic cold sensitivity in the reported range. Another important question is how TRPA1 responses compare to sensory neural responses to cold. Cold responses in dissociated sensory neurons are heterogeneous.26 Cold-sensitive neurons in rats and mice are divided into at least two populations, based on their pharmacology and their temperature-response proﬁles. The major cold-sensitive population is activated by moderate cooling and is sensitive to the cooling agent menthol, indicating that the cold/menthol receptor TRPM8 contributes to the observed responses. A smaller percentage of sensory neurons respond to cold, but not to menthol, indicating that mechanisms independent from TRPM8 are involved. These could include the activation of other depolarizing conductances such as TRPA1 or the inhibition of hyperpolarizing potassium channels. Comparison of the cellular distribution of TRPA1 with the prevalence of nonmenthol cold-responsive cells has not helped to resolve whether or not TRPA1 is involved in sensing cold. While initial in situ hybridization experiments detected TRPA1 in only 3.6 percent of mouse DRG neurons,15 a number comparable to the prevalence of nonmenthol cold-responsive cells, consensus is building that TRPA1 expression is more widespread. Recent reports found that TRPA1 is expressed in 20–36.7 percent of trigeminal neurons, 20–56.5 percent of DRG neurons, and 28.4 percent of neurons in nodose ganglia.12,17,18,27,28 While these numbers compare very well with the percentages of mustard oil– sensitive cells, they are much larger than the percentage of nonmenthol coldresponsive neurons. Because no signiﬁcant correlation between mustard oil responses and cold sensitivity was found in cultured neurons, Babes et al. concluded that TRPA1 is not essential for the cold response.26 These authors also observed signiﬁcant kinetic differences in the cold responses of menthol-insensitive neurons and of TRPA1 in heterologous cells. Additional comparisons of

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cellular responses to TRPA1 activators such as allicin or cinnamaldehyde with cold responses again led to divergent conclusions about the role of TRPA1 in cold sensation.12,18,19 Cellular and behavioral analyses of TRPA1-deﬁcient mice showed that TRPA1 is not essential for acute responses to cold.20 The prevalence of menthol-insensitive cold-responsive neurons remained unchanged when compared with wild-type mice, indicating that a TRPA1-independent mechanism is responsible for activation of these cells by cold. Behavioral responses to evaporative cooling and to different temperatures on the cold plate were completely normal in TRPA1-deﬁcient mice.20 Thus, TRPA1 is not required for cold-induced pain in vivo. As noxious cold sensitivity of sensory neurons does not depend on TRPA1 expression, we are left with the question of whether the cold-activated currents in TRPA1-expressing heterologous cells that have been observed by some investigators can be attributed to intrinsic cold sensitivity of the channel or whether such currents are the result of indirect mechanisms, channel overexpression, disturbances of intracellular Ca2+ levels, or other causes. While TRPA1 is not essential for acute cold-induced pain, a recent report found that the suppression of TRPA1 by antisense nucleotides reduces hypersensitivity to cold temperature (cold allodynia) in rat models of inﬂammation and nerve injury.27 These authors also found that the transcription of TRPA1 is increased during inﬂammation. These data suggest that TRPA1 expression and regulation may affect the excitability of temperature-sensitive neurons in vivo.

TRPA1 IS A RECEPTOR-OPERATED CHANNEL ACTING IN CONCERT WITH TRPV1 TRP ion channels are regulated through signaling pathways that are activated by G-protein-coupled receptors and other membrane receptors. In sensory neurons, TRPV1 activity is regulated by a number of phospholipase C (PLC)–coupled receptors, including the B2 bradykinin receptor, TrkA, the receptor for nerve growth factor (NGF), and ATP receptors. Bradykinin and NGF sensitize TRPV1 to heat and to noxious chemical stimuli, leading to thermal hyperalgesia. Heat sensitivity of TRPV1 is increased by PLC-mediated pathways that remove inhibition of TRPV1 by the membrane phospholipid PIP2.29,30 Heterologous cells coexpressing TRPA1 and PLC-coupled receptors respond with immediately activating cationic currents when receptor agonists are applied.12,16 This indicates that TRPA1 may function as a receptor-operated channel in vivo. Bradykinin, in addition to causing thermal hyperalgesia through sensitization of TRPV1, elicits acute pain through immediate depolarization of sensory neurons. This effect, while diminished, is maintained in TRPV1-deﬁcient mice, suggesting that additional conductances such as TRPA1 may be required for full neural excitation. Indeed, neurons isolated from TRPA1-deﬁcient mice fail to display bradykinin-induced Ca2+ inﬂux.20 Even more strikingly, bradykinin-induced thermal hyperalgesia is absent in TRPA1-deﬁcient mice. While thermal hyperalgesia was initially attributed solely to TRPV1, this result indicates that TRPA1 and TRPV1 are interdependently regulated downstream of BK2 receptors to establish hypersensitivity to heat. Apparently, TRP channels from different branches of the TRP channel gene family can act in concert

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to exert physiological effects. Whether this interdependence reﬂects direct physical interaction between channels and receptors or an indirect regulatory relationship remains to be investigated.

TRPA1 AS A MULTIPLE IRRITANT SENSOR TRPA1 channels respond to a multitude of irritants with diverse origins and chemical structures. Subsets of sensory neurons in the lung, eyes, and mucous membranes have a broad sensitivity to diverse volatile chemical irritants and environmental toxicants. These compounds induce pain, coughing, apnea, and lachrymation and inspire an individual’s behavior to be protected from further exposure. Responses to most chemical irritants are retained in mice deﬁcient in the capsaicin receptor TRPV1, implicating other mechanisms of neural activation. Is TRPA1 involved as a multiple irritant sensor in these neurons? How is such a broad range of sensitivity achieved? Currently, it is unknown whether or not mustard oil and other activators interact with TRPA1 directly. We can discuss different options for the mechanism of channel activation. First, chemical irritants may bind to TRPA1 through “classical” ligand-receptor interaction. While most receptor-ligand systems require high degrees of speciﬁcity, some receptors bind multiple ligands with diverse structures. For example, bitter-taste receptors are known to interact with a multitude of plant alkaloids and other bitter-tasting chemicals that don’t show much resemblance to each other in their chemical structures.31 Similarly, TRPA1 may bind to a large variety of irritant molecules to induce sensory neural excitation. Second, chemically reactive irritants such as mustard oil could form permanent or transient covalent bonds with TRPA1, thereby activating the channel. Isothiocyanates, allicin, and unsaturated aldehydes are reactive compounds capable of forming covalent bonds with cysteine and other residues in proteins. Third, reactive irritants could interfere with signaling pathways that regulate TRPA1, leading to channel activation. These pathways could include phosphorylation cascades, or regulation of intracellular Ca2+ that is known to affect TRPA1 function.12

CONCLUSION The studies discussed in this chapter show that TRPA1 channels in mammalian sensory neurons contribute to acute and inﬂammatory pain. TRPA1 is activated downstream of inﬂammatory PLC-coupled receptors for bradykinin and other proalgesic agents in vivo and acts in concert with TRPV1 to cause thermal hyperalgesia. Whether endogenous ligands for TRPA1 exist remains to be established. The sensitivity of TRPA1 to phytocannabinoids indicates that endogenous cannabinoid-like ligands may modulate TRPA1 function. While it is not known whether mustard oil–like compounds are endogenously expressed in mammals, other endogenous organosulfur compounds could affect the TRPA1 activity. Future pharmacological, electrophysiological, and genetic studies are needed to clarify these and other aspects of TRPA1 function. In addition to TRPV1, TRPA1 represents an exciting new target for the development of potential new analgesics and anti-inﬂammatory agents.

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REFERENCES 1. Bruce, A.N., Über die Beziehungen der sensiblen Nervenendigungen zum Entzündungsvorgang, Arch. Exp. Pathol. Pharmakol. 63, 423–33, 1910. 2. Louis, S.M., Johnstone, D., Russell, N.J., Jamieson, A., and Dockray, G.J., Antibodies to calcitonin-gene related peptide reduce inﬂammation induced by topical mustard oil but not that due to carrageenin in the rat, Neurosci. Lett. 102 (2–3), 257–60, 1989. 3. Smith, C.H., Barker, J.N., Morris, R.W., MacDonald, D.M., and Lee, T.H., Neuropeptides induce rapid expression of endothelial cell adhesion molecules and elicit granulocytic inﬁltration in human skin, J. Immunol. 151 (6), 3274–82, 1993. 4. Jancso, N., Jancso-Gabor, A., and Szolcsanyi, J., Direct evidence for neurogenic inﬂammation and its prevention by denervation and by pretreatment with capsaicin, Br. J. Pharmacol. 31 (1), 138–51, 1967. 5. Simons, C.T., Carstens, M.I., and Carstens, E., Oral irritation by mustard oil: selfdesensitization and cross-desensitization with capsaicin, Chem. Senses 28 (6), 459–65, 2003. 6. Gamse, R., Holzer, P., and Lembeck, F., Decrease of substance P in primary afferent neurones and impairment of neurogenic plasma extravasation by capsaicin, Br. J. Pharmacol. 68 (2), 207–13, 1980. 7. Cuello, A.C., Gamse, R., Holzer, P., and Lembeck, F., Substance P immunoreactive neurons following neonatal administration of capsaicin, Naunyn Schmiedebergs Arch. Pharmacol. 315 (3), 185–94, 1981. 8. Hori, T. and Tsuzuki, S., Thermoregulation in adult rats which have been treated with capsaicin as neonates, Pﬂügers Arch. 390 (3), 219–23, 1981. 9. Caterina, M.J., Schumacher, M.A., Tominaga, M., Rosen, T.A., Levine, J.D., and Julius, D., The capsaicin receptor: a heat-activated ion channel in the pain pathway, Nature 389 (6653), 816–24, 1997. 10. Caterina, M.J., Lefﬂer, A., Malmberg, A.B., Martin, W.J., Trafton, J., Petersen-Zeitz, K.R., Koltzenburg, M., Basbaum, A.I., and Julius, D., Impaired nociception and pain sensation in mice lacking the capsaicin receptor, Science 288 (5464), 306–13, 2000. 11. Davis, J.B., Gray, J., Gunthorpe, M.J., Hatcher, J.P., Davey, P.T., Overend, P., Harries, M.H., Latcham, J., Clapham, C., Atkinson, K., Hughes, S.A., Rance, K., Grau, E., Harper, A.J., Pugh, P.L., Rogers, D.C., Bingham, S., Randall, A., and Sheardown, S.A., Vanilloid receptor-1 is essential for inﬂammatory thermal hyperalgesia, Nature 405 (6783), 183–87, 2000. 12. Jordt, S.E., Bautista, D.M., Chuang, H.H., McKemy, D.D., Zygmunt, P.M., Hogestatt, E.D., Meng, I.D., and Julius, D., Mustard oils and cannabinoids excite sensory nerve ﬁbres through the TRP channel ANKTM1, Nature 427 (6971), 260–65, 2004. 13. Patacchini, R., Maggi, C.A., and Meli, A., Capsaicin-like activity of some natural pungent substances on peripheral endings of visceral primary afferents, Naunyn Schmiedebergs Arch. Pharmacol. 342 (1), 72–77, 1990. 14. Wilson, R.K., Kwan, T.K., Kwan, C.Y., and Sorger, G.J., Effects of papaya seed extract and benzyl isothiocyanate on vascular contraction, Life Sci. 71 (5), 497–507, 2002. 15. Story, G.M., Peier, A.M., Reeve, A.J., Eid, S.R., Mosbacher, J., Hricik, T.R., Earley, T.J., Hergarden, A.C., Andersson, D.A., Hwang, S.W., McIntyre, P., Jegla, T., Bevan, S., and Patapoutian, A., ANKTM1, a TRP-like channel expressed in nociceptive neurons, is activated by cold temperatures, Cell 112 (6), 819–29, 2003.

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16. Bandell, M., Story, G.M., Hwang, S.W., Viswanath, V., Eid, S.R., Petrus, M.J., Earley, T.J., and Patapoutian, A., Noxious cold ion channel TRPA1 is activated by pungent compounds and bradykinin, Neuron 41 (6), 849–57, 2004. 17. Nagata, K., Duggan, A., Kumar, G., and Garcia-Añoveros, J., Nociceptor and hair cell transducer properties of TRPA1, a channel for pain and hearing, J. Neurosci. 25 (16), 4052–61, 2005. 18. Bautista, D.M., Movahed, P., Hinman, A., Axelsson, H.E., Sterner, O., Hogestatt, E.D., Julius, D., Jordt, S.E., and Zygmunt, P.M., Pungent products from garlic activate the sensory ion channel TRPA1, Proc. Natl. Acad. Sci. USA 102 (34), 12248–52, 2005. 19. Macpherson, L.J., Geierstanger, B.H., Viswanath, V., Bandell, M., Eid, S.R., Hwang, S., and Patapoutian, A., The pungency of garlic: activation of TRPA1 and TRPV1 in response to allicin, Curr. Biol. 15 (10), 929–34, 2005. 20. Bautista, D.M., Jordt, S.E., Nikai, T., Tsuruda, P.R., Read, A.J., Poblete, J., Yamoah, E.N., Basbaum, A.I., and Julius, D., TRPA1 mediates the inﬂammatory actions of environmental irritants and proalgesic agents, Cell 124 (6), 1269–82, 2006. 21. Jones, R.T., Cardiovascular system effects of marijuana, J. Clin. Pharmacol. 42 (11 Suppl.), 58S–63S, 2002. 22. Pacher, P., Batkai, S., and Kunos, G., Blood pressure regulation by endocannabinoids and their receptors, Neuropharmacology 48 (8), 1130–38, 2005. 23. Hogestatt, E.D. and Zygmunt, P.M., Cardiovascular pharmacology of anandamide, Prostaglandins Leukot. Essent. Fatty Acids 66 (2–3), 343–51, 2002. 24. Zygmunt, P.M., Petersson, J., Andersson, D.A., Chuang, H., Sørgård, M., Di Marzo, V., Julius, D., and Högestätt, E.D., Vanilloid receptors on sensory nerves mediate the vasodilator action of anandamide, Nature 400 (6743), 452–57, 1999. 25. Zygmunt, P.M., Andersson, D.A., and Hogestatt, E.D., Delta 9-tetrahydrocannabinol and cannabinol activate capsaicin-sensitive sensory nerves via a CB1 and CB2 cannabinoid receptor-independent mechanism, J. Neurosci. 22 (11), 4720–27, 2002. 26. Babes, A., Zorzon, D., and Reid, G., Two populations of cold-sensitive neurons in rat dorsal root ganglia and their modulation by nerve growth factor, Eur. J. Neurosci. 20 (9), 2276–82, 2004. 27. Obata, K., Katsura, H., Mizushima, T., Yamanaka, H., Kobayashi, K., Dai, Y., Fukuoka, T., Tokunaga, A., Tominaga, M., and Noguchi, K., TRPA1 induced in sensory neurons contributes to cold hyperalgesia after inﬂammation and nerve injury, J. Clin. Invest. 115 (9), 2393–401, 2005. 28. Kobayashi, K., Fukuoka, T., Obata, K., Yamanaka, H., Dai, Y., Tokunaga, A., and Noguchi, K., Distinct expression of TRPM8, TRPA1, and TRPV1 mRNAs in rat primary afferent neurons with adelta/c-ﬁbers and colocalization with trk receptors, J. Comp. Neurol. 493 (4), 596–606, 2005. 29. Chuang, H.H., Prescott, E.D., Kong, H., Shields, S., Jordt, S.E., Basbaum, A.I., Chao, M.V., and Julius, D., Bradykinin and nerve growth factor release the capsaicin receptor from PtdIns(4,5)P2-mediated inhibition, Nature 411 (6840), 957–62, 2001. 30. Prescott, E.D. and Julius, D., A modular PIP2 binding site as a determinant of capsaicin receptor sensitivity, Science 300 (5623), 1284–88, 2003. 31. Mueller, K.L., Hoon, M.A., Erlenbach, I., Chandrashekar, J., Zuker, C.S., and Ryba, N.J., The receptors and coding logic for bitter taste, Nature 434 (7030), 225–29, 2005.

12

TRPA1 in Auditory and Nociceptive Organs Jaime García-Añoveros and Anne Duggan Northwestern University

CONTENTS Distribution of TRPA1 in the Inner Ear................................................................164 Functional Tests of TRPA1 in the Inner Ear ........................................................165 Comparing Channel Properties of TRPA1 and of the Hair Cell Transducer.......166 Distribution of TRPA1 in Sensory Ganglia ..........................................................169 The Function of TRPA1 in Nociceptors ...............................................................170 A Channel Property of TRPA1 for Nociception...................................................172 TRPA1 in Invertebrates .........................................................................................172 Note........................................................................................................................173 References..............................................................................................................173 TRPA1, initially called P120 and ANKTM1, was originally described as a downregulated protein in mesenchymal tumor cells and was detected in cultured ﬁbroblasts but lost upon oncogenic transformation, although no expression in healthy tissues was described (Schenker and Trueb, 1998; Jaquemar et al., 1999). With in situ hibridization, we found mouse TRPA1 mRNA absent from most major organs (brain, heart, liver, kidneys, skeletal muscles, lungs, spleen, and testes, as well as whisker pad skin and superior cervical ganglia), but present in the inner ear and in certain peripheral sensory ganglia: dorsal root (DRG), trigeminal (TG), and nodose (Nagata et al., 2005). This very restricted pattern of expression suggests speciﬁc roles unique to sensory function. Molecularly, TRPA1 has six membrane-spanning domains and a presumed poreforming domain characteristic of all TRPs and many other ion channels. Its N- and C-terminal segments are predicted to be cytoplasmic. In addition, a distinguishing feature of TRPA1 is a very long N-terminus with up to 17 predicted ankyrin (ANK) repeats (Figure 12.1). Many similar repeats are present in the protein ankyrin, and 29 of them are present in TRPN1, a mechanosensory channel protein of ﬂies, worms, and ﬁsh that has not been found in mammals (Walker et al., 2000; Sidi et al., 2003; Li et al., 2006). ANK repeats may serve at least two functions: (1) to interact with other proteins, particularly those of the cytoskeleton, and (2) to provide elasticity when in tandem, forming a molecular spring (cited as V. Bennet, personal

163

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FIGURE 12.1 Domain structure of TRPA1 with its predicted topology (the lipid membrane is represented by a gray band) and phylogenetic comparison with other TRP channels of mammals (bold) and nematodes (italic). Because there is no mammalian NOMPC (TRPN1), we include the insect (bold-italic) and zebraﬁsh (underline) orthologues. Mammalian TRPA1 has a putative orthologue in C. elegans protein C29E6.2 (hence renamed TRPA1).

communication, in Corey et al., 2004; Howard and Bechstedt, 2004; Sotomayor et al., 2005; Lee et al., 2006). Because many models of mechanically gated channels postulate linkage to a force-transducing structure like the cytoskeleton or an elastic gating spring (García-Añoveros and Corey, 1997; Gillespie and Walker, 2001), channels like TRPN1 and TRPA1 may be well endowed for the mechanical transduction that characterizes the auditory and vestibular organs as well as the somatosensory and autonomic ganglia that express it. Based on the hypothesis that the transduction channel of hair cells and somatosensory neurons could be a TRP channel (Duggan et al., 2000), we screened (RTPCR and in situ hybridization) TRP genes for expression in the inner ear and somatosensory ganglia. We found several TRPs expressed in ganglia and two whose mRNA is expressed in the organ of Corti, the place in the cochlea where the sensory cells reside (unpublished results and Corey et al., 2004). In addition, a third TRP channel protein (MCOLN3, or TRPML3) has also been detected in hair cells (Di Palma et al., 2002). In this chapter we discuss TRPA1, which is expressed both in the inner ear and peripheral ganglia.

DISTRIBUTION OF TRPA1 IN THE INNER EAR By ISH we found TRPA1 expression in supporting cells of the neonatal organ of Corti, although there was no clearly detectable expression in the mechanosensory hair cells (Corey et al., 2004). However, because very few functional transduction channels are present per hair cell, very low levels of TRPA1 are not inconsistent

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with this protein contributing to transduction channels. We ﬁrst considered this exciting, if less straightforward, possibility. Using an antibody to the N-terminus of TRPA1, we conﬁrmed expression of this protein in all the mechanosensory epithelia of the inner ear: cochlear organ of Corti, saccular and utricular maculae, and crista ampullaris of the semicircular canals. The immunoreactivities were completed by excess antigenic peptide. Further, this antibody recognizes a single band in Western blots from organ of Corti or saccule plus utricle. Therefore our antibody appears to speciﬁcally recognize TRPA1. These experiments conﬁrmed strong TRPA1 expression in supporting cells, but also revealed a late onset TRPA1 immunoreactivity to the sensory hair cells at the kinocilia, structures that are distinct from the actin-rich mechanosensory stereocilia and are not thought to be mechanosensory, but also at cuticular plates and, weakly, the stereocilia (Kumar et al., 2005; Nagata et al., 2005). Studies using an antibody raised against the C-terminus, although not as extensively controlled, report similar immunoreactivity (Corey et al., 2004). However, stereocilia have been known for nonspeciﬁcally binding antibodies. Further experiments are needed to conﬁrm functional TRPA1 presence in hair cells. In support of this, TRPA1 agonists AITC and icilin were recently reported to activate inward currents in hair cells (Stepanyan et al., 2006). Because mechanosensory transduction takes place at the stereocilia from day 17 of mouse embryogenesis, and the cuticular plate is thought to hold a reserve of transduction components, this localization pattern is consistent with a potential role of TRPA1 in hair cell mechanotransduction, but also suggests that other proteins make transduction channels early in development.

FUNCTIONAL TESTS OF TRPA1 IN THE INNER EAR Two general approaches address the function of TRPA1 in hearing: (1) inhibiting its production with acutely applied agents (Corey et al., 2004), and (2) mutating the gene by deleting part of it (Bautista et al., 2006). These two approaches have yielded apparently contradictory results. TRPA1 knockdown with morpholinos in zebraﬁsh or with viral-mediated RNA interference in cultured mice utricles resulted in a reduction, although not an elimination, in the magnitude of the hair cell transduction currents. Although these agents can have nonspeciﬁc effects, an alternative interpretation is that they speciﬁcally inhibited TRPA1 production and that this protein participates, directly or indirectly, in generating mechanotransduction currents. On the other hand, a deletion of the pore domain of TRPA1-produced animals with apparently normal auditory function, as assessed by auditory brainstem responses (ABRs) and distortion product otoacustic emissions (DPOAEs), clearly indicates that intact TRPA1 is not necessary for hair cell transduction. However, we should bear in mind that (1) the deletion of TRPA1 left a large portion of the protein intact (the 17 ANK repeats plus the ﬁrst ﬁve membrane-spanning domains), and (2) TRPA1 may act redundantly with other channel subunits to form transduction channels. The fact that there are transduction channels distributed along the length of the cochlea with

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different conductance levels (Ricci et al., 2003; He et al., 2004) suggests to us that these channels are molecularly heterogeneous, something that could be accomplished by alternative splicing or by heteromultimerization of several distinct channel subunits (like other TRPs that we and others have found in hair cells). In this admittedly hypothetical context, a protein like TRPA1 could contribute to hair cell transducers but be compensated for by other subunits if lacking, perhaps accounting for the lack of phenotype of the mutant and the limited effects of the acute inhibition studies. Although the mutant phenotype proves that TRPA1 is not an essential subunit of the pore, we still need to determine with certainty whether TRPA1 contributes to hair cell transduction or not.

COMPARING CHANNEL PROPERTIES OF TRPA1 AND OF THE HAIR CELL TRANSDUCER If TRPA1 is indeed a component of the hair cell transducer, we would expect that the pore properties of TRPA1 in heterologous cells and those of the hair cell transducer to be similar, with some distinctions due to the lack of other subunits in heterologous cells. Heterologously expressed TRPA1 can be opened with several agonists such as allyl isothiocyanate (AITC) (Bandell et al., 2004; Jordt et al., 2004; Nagata et al., 2005). Hence, TRPA1 channel properties could be studied and compared with those already known for the endogenous hair cell mechanotransducing channel. Many similarities, and a few differences, have been found (Nagata et al., 2005). The most prominent difference relates to channel gating. Heterologously expressed TRPA1 has not been shown to be mechanically gated. This is not surprising: in hair cells the channel is in a unique subcellular specialization, the stereocilia, with a unique molecular composition of lipids and proteins. Indeed, hair cell stereocilia are rich in certain lipids like PIP2, and its removal impairs mechanotransduction (Hirono et al., 2004). Most models of hair cell transduction postulate that the channel associates with other proteins, like the extracellular tip link and a linker to the actin cytoskeleton, which transmit gating force to the channel. Without these structures, a channel may not be able to sense displacements. Despite these differences in gates in distinct cellular settings, if TRPA1 forms part of the hair cell transducer one would expect that, once opened, the properties of their pores would be similar, as it is the pore that TRPA1 would contribute to in the hair cell transduction complex. Heterologous TRPA1 (Nagata et al., 2005) and the hair cell transducer (Jorgensen and Ohmori, 1988; Kroese et al., 1989; Rüsch et al., 1994; Kimitsuki et al., 1996; Ricci, 2002; Farris et al., 2004) are blocked by the same four antagonists (amiloride, gadolinium, gentamicin, and ruthenium red), and all with indistinguishable Hill coefﬁcients, which supports a common mode of action. Of these blockers, the two that act by plugging into the pore, ruthenium red and gentamicin, have IC50s that are indistinguishable between TRPA1 in heterologous cells and the mechanotransducer in hair cells, but the two other blockers act with different afﬁnities. Heterologously expressed TRPA1 is 100 times more sensitive to Gd3+ (it is indeed the most gadolinium-sensitive channel known) and 10 times less sensitive

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to amiloride, revealing some molecular divergence between heterologous TRPA1 and endogenous hair cell transducers. Another parallel between TRPA1 and the hair cell transducer relates to the effects of calcium on channel gating and conductance. Calcium, a permeant ion of both channels, reduces single channel conductance to the same extent (54 percent of its conductance with no or just micromolar amounts of Ca2+) (Ricci et al., 2003; Nagata et al., 2005). In addition, as extracellular Ca2+ enters the transduction channel, it binds to a site at or very close to the channel and causes ﬁrst an increase in open probability (as detected with single channel recordings in hair cells) and then closure, a phenomenon often referred to as fast adaptation (Howard and Hudspeth, 1988; Kennedy et al., 2003). Upon depolarization, the channels open again (Ricci et al., 2000). The same events take place in heterologously expressed TRPA1 chemically activated with AITC (Nagata et al., 2005): (1) Ca2+ entering from the outside causes (as mentioned above) a reduction of single channel conductance, but also increases the open probability, resulting in a brief potentiation of the current. (2) Subsequently, Ca2+ induces channel closure (Figure 12.2a, b, and c). It should be noted that, while the Ca2+-induced closure is thought to mediate fast adaptation in hair cells, the brief potentiation of opening might underlie the “release” that occurs just before adaptation, which fosters a small, additional positive displacement of the hair bundle, a phenomenon that may contribute to ampliﬁcation of the response of the cochlea, and thus to enhancing hearing sensitivity to soft sounds (Hudspeth, 2005; Lemasurier and Gillespie, 2005) (Figure 12.2d). (3) Through an unknown mechanism, cellular depolarization after calciuminduced closure reopens the channels. Figure 12.2c shows a model of calcium action, modiﬁed from one developed to account for the behavior of the hair cell transducer but that also illustrates the phenomena described for heterologously expressed TRPA1. The primary caveats to this comparison are due to timing, for while the effects of Ca2+ on the hair cell transducer take place in milliseconds, in chemically activated, heterologously expressed TRPA1 they take seconds. However, these quantitative differences do not imply a mechanistic difference. What we envision might be happening is that Ca2+, as it goes through the channel, either goes across to the cytoplasm or binds to it, in which case it induces ﬁrst potentiation (by increasing Po, even if single channel conductance diminishes), and, second, closure (Figure 12.2c). Most calcium ions go through, and how long it takes for one to bind would depend on the afﬁnity of the binding site, something that could vary between TRPA1 expressed alone or as part of the transduction complex in hair cells. Finally, TRPA1 displayed a single channel conductance of ~100 pS (Nagata et al., 2005), similar to that described for the hair cell transducer (Crawford et al., 1991; Denk et al., 1995; Géléoc et al., 1997; Ricci et al., 2003). Given the many mechanistic similarities between the pores of TRPA1 and the hair cell transducers, but also considering the various quantitative differences, it

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FIGURE 12.2 Effects of calcium on permeation and gating of chemically activated (with AITC), heterologously expressed TRPA1 shown for (a) single channels (in outside out patches) and (b) whole cells (Nagata et al., 2005). (c) Calcium effects on TRPA1 as well as on the hair cell transducer can both be described with the same model (modiﬁed from Farris et al., 2004). (d) We propose that these channel gating phenomena may account for some of the mechanical properties of transducing hair cell bundles (modiﬁed from Lemasurier and Gillespie, 2005). Roman numerals indicate equivalent states in single channel recordings, whole-cell recordings, and theoretical models.

seems that either TRPA1 or similar channel proteins (perhaps the other TRPs found in stereocilia), or perhaps heteromultimers of TRPA1 with these various proteins, could form the pore of the hair cell mechanotransducer. At this point, more experimentation is required to determine the function of TRPA1 in hair cells, especially whether it does or does not contribute to mechanotransducing channels. Yet another intriguing question is what the function of TRPA1 might be

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not in hair cells but in their support cells, where its expression, although not yet thoroughly described, is most prominent.

DISTRIBUTION OF TRPA1 IN SENSORY GANGLIA Outside the ear, TRPA1 mRNA has only been reproducibly detected in peripheral sensory ganglia that have nociceptive neurons: trigeminal, dorsal root, and nodose. In all these ganglia, the nociceptive neurons are generally the smaller cells in diameter and axon caliber. In situ hybridization in mouse sections demonstrated TRPA1 mRNA expression by a large number of the smaller, nociceptive cells (36.5 percent of all neurons in TG, 56.5 percent of all neurons in cervical DRGs, and 28.4 percent of all neurons in nodose) (Nagata et al., 2005). Similar results were obtained in rats (36.7 percent in TG, 39.5 percent in lumbar, L5 and L6 DRGs) and indicated that these cells do not coexpress neuroﬁlament 200 or the growth factor receptors TrkC and TrkB, but do express the NGF receptor TrkA or no Trk receptor at all; most express other markers of nociceptive neurons like the capsaicin receptor TRPV1, calcitonin gene-related peptide (CGRP), or substance P (SP) (Kobayashi et al., 2005). All these experiments conﬁrm that TRPA1 is expressed by C-ﬁber nociceptors and not by A-ﬁber neurons (including the Aδ ﬁbers, the faster-conducting nociceptors, as well as the Aβ and Aα innocuous mechanoreceptors). A previous report indicating TRPA1 expression in only 3.6 percent of the mouse DRG neurons may be an underestimate (Story et al., 2003), whereas another report of expression based on neonatal rat TG (~20%) (Jordt et al., 2004) is similar enough given the potential for developmental differences. However, it is important to realize that experimentally induced inﬂammation and neuropathic pain increase the number of neurons that express TRPA1 mRNA (Obata et al., 2005), so the precise distribution may depend on the nociceptive history of each animal and ganglion. As a general rule, however, TRPA1 is expressed by half or more of the small, C-ﬁber nociceptive neurons, a distribution consistent with a prominent role for TRPA1 channels in pain. Antibodies to TRPA1 conﬁrm expression to small, peripherin-positive nociceptors (Bautista et al., 2005; Nagata et al., 2005). In addition to detecting TRPA1 protein at the cell bodies (the site of protein synthesis), it is also detected in their peripheral nerves, the expected subcellular location for a channel involved in nociceptive transduction. Unlike many other potential mediators of pain, TRPA1 has a surprisingly restricted expression, having been so far only reproducibly detected in a few cells of the inner ear and in most nociceptors. Therefore, inhibitors of this channel, provided that they are speciﬁc (currently known ones are not), could perhaps function as pain-killers with limited side effects. Delivery to the ear may have to be avoided, but this would be easily achieved with local applications, which should be effective because TRPA1 is present at the periphery. A drug with these characteristics could revolutionize the medical treatment of pain. But what kind of pain would a TRPA1 antagonist block?

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THE FUNCTION OF TRPA1 IN NOCICEPTORS Functional evidence that TRPA1 may indeed serve as a pain-receptor channel ﬁrst came from heterologous expression studies that demonstrate that TRPA1 channels are opened upon exposure to various pain-producing chemicals (the pungent components of many often edible substances): allyl isothiocyanate (horseradish, wasabi, and mustard oil), benzil isothiocyanate (yellow mustard and crushed papaya seeds), phenylethyl isothiocyanate (Brussels sprouts), methyl isothiocyanate (capers and nasturtium seeds), cinnamaldehyde (cinnamon), methyl salicilate (wintergreen oil, often used in mouthwashes like Listerine), eugenol (clove), allicin (freshly crushed garlic), the synthetic compound AG-3-5 (icilin), and acrolein (tear gas and the undesirable by-product of certain chemotherapeutic regimes) (Story et al., 2003; Bandell et al., 2004; Jordt et al., 2004; Bautista et al., 2005; Macpherson et al., 2005; Nagata et al., 2005; Bautista et al., 2006). More recently, mice with a deletion of part of TRPA1 have been generated and found to be insensitive to TRPA1 agonists mustard oil and allicin. Dissociated trigeminal ganglion neurons from mutants did not respond to these agonists, and mutant animals did not react adversely to their acute application and did not develop neurogenic inﬂammation and hyperalgesia, which wild-type animals do. This suggests that TRPA1 is the primary and perhaps only receptor for these pungent compounds in nociceptive neurons (Bautista et al., 2006). How all these different compounds activate TRPA1 is not clear. Their low hydrophilicity and extremely slow activation (tens of seconds to minutes), plus the fact that they can activate a cell-attached patch by exposure to the outside of the rest of the cell membrane, imply a mechanism of action that might require prior partition of these chemicals into the lipid membrane to then, directly or indirectly, activate TRPA1. Because these pungent chemicals are not endogenous to animals and no evidence indicates that they would appear in damaged tissues, they would not, under normal physiological conditions, activate TRPA1. What does? Another mechanism that opens TRPA1 in heterologous cells is through metabotropic receptors that activate phospholipase C (PLC) second-messenger pathways (Jordt et al., 2004). This is interesting because many endogenous pro-algesics and pro-inﬂammatory agents (bradykinin, histamine, serotonin, ATP, and neurotrophins) act this way. Indeed, bradykinin has been shown to activate TRPA1 in cells that also expressed the bradykinin receptor (Bandell et al., 2004). Accordingly, TRPA1 mutant mice did not develop hyperalgesia in response to bradykinin. Interestingly, for responding to bradykinin, trigeminal neurons require both TRPA1 and TRPV1 (Bautista et al., 2006). Hence TRPV1, which is an ionotropic sensor for noxious heat and acidosis, can also act as a mediator of second messenger–induced sensitization. We wonder if there may also be an ionotropic noxious stimulus, besides the above-mentioned exogenous agonists, that activates TRPA1. An interesting hypothesis is that TRPA1 is a sensor for painfully cold temperatures. This idea was suggested by heterologous expression studies of TRPA1 that showed channel activation by temperatures in the painfully cold range (below 17°C) and by icilin (Story et al., 2003; Bandell et al., 2004), a synthetic compound that produces a cool sensation. A major drawback to this hypothesis is that, in attempts

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by others, heterologously expressed TRPA1 has been activated by icilin, but not by cold (Jordt et al., 2004; Nagata et al., 2005). In addition, TRPA1 mutant mice respond normally to cold, and their sensory ganglia have a normal distribution of mentholsensitive (i.e., TRPM8 expressing) and insensitive neurons (Bautista et al., 2006). Because icilin also activates the cold receptor channel TRPM8, and it produces both a cold and a prickling sensation, it may be that the cold sensation is mediated by TRPM8 activation and the prickling by TRPA1 activation. Perhaps in certain cellular settings cold triggers a cellular response that can activate TRPA1, which is not intrinsically sensitive to cold itself, thus explaining the occasional currents observed upon cooling in cells expressing TRPA1. But whether TRPA1 plays a physiological role in sensing painful cold seems at present unlikely (Reid, 2005). In addition, all other chemical agonists of TRPA1 produce subjective sensations of pain, but not of cold, arguing against a role of TRPA1 in sensing cold. It is certainly possible that painful cold sensation results from the combined activation of TRPA1 and TRPM8, whereas pungent chemicals that activate TRPA1 do not act on TRPM8 and could thus produce a different pain. But even this would imply that TRPA1 mediates something else than painful cold. One hypothesis is force. Regardless of the function of TRPA1 in the ear, its pharmacology indicates that it is blocked by a set of four chemicals (amiloride, Gd3+, ruthenium red, and gentamycin) that are characteristic blockers of mechanosensory channels found in several cell types besides hair cells (Hamill and McBride, 1996; Nagata et al., 2005). The wide distribution of TRPA1 among nociceptors is consistent with the large number of c-ﬁbers that are or can become mechanosensitive. A caveat to this hypothesis is that in heterologous cells, TRPA1 has not thus far been shown to be mechanically gated, although this may be explained by the lack of accessory molecules, both proteins and lipids, that would confer mechanosensitivity to a channel. Perhaps more important is that TRPA1 mutant mice have the same paw withdrawal thresholds to mechanical stimulation as wild-type mice (Bautista et al., 2006). However, this is not necessarily unexpected if we consider the lack of naturally occurring mutants, either in mice or in humans, with congenital insensitivity to touch or mechanically induced nociception. This rarity, striking in comparison with the hundreds of genes that, when mutated, produce other sensory defects like deafness or blindness, suggests that either (1) very few genes participate in somatosensory mechanoreception, which seems unlikely, or (2) many genes participate, and do so redundantly. Therefore determination of TRPA1’s potential role in mechanonociception may require detailed phenotypic analysis or the generation of animals with mutations in other genes in addition to TRPA1. TRPA1 may very well be polymodal, activated by more than one noxious form of stimulation, as other nociceptive receptor channels have been found to be (for example, the capsaicin-, heat-, and acid-sensitive TRPV1, which also mediates bradykinin-induced sensitization). In this regard, it is worth noting that in Drosophila the painless gene, which encodes a TRPA homologue (not the TRPA1 orthologue, but a close relative with no mammalian orthologue), mediates the aversive response to touch with a heated probe and is required for both thermal and mechanical nociception (Tracey et al., 2003).

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A CHANNEL PROPERTY OF TRPA1 FOR NOCICEPTION Whatever the stimulus that gates TRPA1 in nociceptors may be, one property of this channel renders it well suited for nociception (Nagata et al., 2005). As described above, after the channel opens in response to a pungent agonist, extracellular calcium enters, and within tens of seconds it causes channel closure. But this phenomenon is voltage sensitive, so that inactivation occurs if the cell is at a potential of –80 mV but not (or very slowly) at –20 mV, which is the potential reached by a stimulated nociceptive DRG neuron (Blair and Bean, 2003). Therefore, if TRPA1 is weakly stimulated and the resulting currents fail to depolarize the neuronal terminal sufﬁciently, the channels would close. But if a stronger stimulation triggers enough cation inﬂux to sufﬁciently depolarize the neuron, the channel would remain open as long as it is stimulated by the noxious agonist. In this way TRPA1 could distinguish between subthreshold (i.e., innocuous) stimulation, to which it would inactivate, and suprathreshold (i.e., noxious) stimulation, to which it would remain open as long as it lasted. This channel property might account for the lack of habituation that is so characteristic of pain sensation. Furthermore, it is easy to envision that depolarization due to activation of another nociceptive channel (i.e., heat activation of TRPV1, which is expressed by the TRPA1expressing neurons) would lower the threshold stimulation required to keep TRPA1 open. This scenario could account for other characteristic pain phenomena like coincidence detection and perhaps also allodinia (if a sensitizing stimulus were to leave the neuron or its terminal depolarized).

TRPA1 IN INVERTEBRATES Another aspect in which TRPA1 is remarkable is its evolutionary conservation. A phylogenetic comparison of TRP channels from mammalian and nematode genomes (Figure 12.1) reveals that both contain a TRPA1 orthologue, closer to each other than to any other TRP channel produced by their respective genomes, and these orthologues have the same overall protein structure (17 ANK repeats followed by the TRP-like channel domains). We have found expression of C. elegans TRPA1 in presumed nociceptive and mechanosensory neurons, as well as in neuronal support cells, implying a conservation of function from nematodes to mammals (Mancillas et al., 2005, and our unpublished results). This suggests that TRPA1 was used by organisms 600 million years ago in ancestral sensory systems. On the other hand, a distinction of TRPA1 in nematodes is that it is also expressed, rather prominently, in various nonneural cells, some of which are essential for survival. Curiously, most of the pungent agonists of TRPA1 kill worms (our unpublished results), and worm parasites are commonly ingested, especially with uncooked foods. This distinction in TRPA1 expression, in sensory cells in humans but also in vital organs in nematodes, might provide a rationale for why so many cuisines from all over the world mix pungent TRPA1 agonists with nutrients, as humans can tolerate and even acquire a “taste” for the same chemicals that kill their parasites.

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NOTE Another targeted mutation of TRPA1 that also deletes the pore domain was recently published (Kwan et al., 2006). A detailed analysis showed that the mutant mice have raised thresholds and decreased withdrawal responses to mechanical stimulation with Von Frey hairs to the paw. These results add support to our hypothesis that, at least in nociceptors, TRPA1 may form mechanosensory channels.

REFERENCES Bandell, M., Story, G.M., Hwang, S.W., Viswanath, V., Eid, S.R., Petrus, M.J., Earley, T.J., and Patapoutian, A. (2004). Noxious cold ion channel TRPA1 is activated by pungent compounds and bradykinin. Neuron 41:849–857. Bautista, D.M., Jordt, S.E., Nikai, T., Tsuruda, P.R., Read, A.J., Poblete, J., Yamoah, E.N., Basbaum, A.I., and Julius, D. (2006). TRPA1 mediates the inﬂammatory actions of environmental irritants and proalgesic agents. Cell 124:1269–1282. Bautista, D.M., Movahed, P., Hinman, A., Axelsson, H.E., Sterner, O., Hèogestèatt, E.D., Julius, D., Jordt, S.E., and Zygmunt, P.M. (2005). Pungent products from garlic activate the sensory ion channel TRPA1. Proceedings of the National Academy of Sciences of the United States of America 102:12248–12252. Blair, N.T. and Bean, B.P. (2003) Role of tetrodotoxin-resistant Na+ current slow inactivation in adaptation of action potential ﬁring in small-diameter dorsal root ganglion neurons. Journal of Neuroscience 23:10338–10350. Corey, D.P., García-Añoveros, J., Holt, J.R., Kwan, K.Y., Lin, S.Y., Vollrath, M.A., Amalﬁtano, A., Cheung, E.L., Derﬂer, B.H., Duggan, A., Géléoc, G.S., Gray, P.A., Hoffman, M.P., Rehm, H.L., Tamasauskas, D., and Zhang, D.S. (2004). TRPA1 is a candidate for the mechanosensitive transduction channel of vertebrate hair cells. Nature 432:723–730. Crawford, A.C., Evans, M.G., and Fettiplace, R. (1991). The actions of calcium on the mechano-electrical transducer current of turtle hair cells. Journal of Physiology 434: 369–398. Denk, W., Holt, J.R., Shepherd, G.M., and Corey, D.P. (1995). Calcium imaging of single stereocilia in hair cells: localization of transduction channels at both ends of tip links. Neuron 15:1311–1321. Di Palma, F., Belyantseva, I.A., Kim, H.J., Vogt, T.F., Kachar, B., and Noben-Trauth, K. (2002). Mutations in Mcoln3 associated with deafness and pigmentation defects in varitint-waddler (Va) mice. Proceedings of the National Academy of Sciences of the United States of America 99:14994–14999. Duggan, A., García-Añoveros, J., and Corey, D.P. (2000). Insect mechanoreception: what a long, strange TRP it's been. Current Biology 10:R384–387. Farris, H.E., LeBlanc, C.L., Goswami, J., and Ricci, A.J. (2004). Probing the pore of the auditory hair cell mechanotransducer channel in turtle. Journal of Physiology 558:769–792. García-Añoveros, J. and Corey, D.P. (1997). The molecules of mechanosensation. Annual Review of Neuroscience 20:567–594. Géléoc, G.S., Lennan, G.W., Richardson, G.P., and Kros, C.J. (1997). A quantitative comparison of mechanoelectrical transduction in vestibular and auditory hair cells of neonatal mice. Proceedings of the Royal Society of London—Series B: Biological Sciences 264:611–621.

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Gillespie, P.G. and Walker, R.G. (2001). Molecular basis of mechanosensory transduction. Nature 413:194–202. Hamill, O.P. and McBride, D.W., Jr. (1996). The pharmacology of mechanogated membrane ion channels. Pharmacological Reviews 48:231–252. He, D.Z., Jia, S., and Dallos, P. (2004). Mechanoelectrical transduction of adult outer hair cells studied in a gerbil hemicochlea. Nature 429:766–770. Hirono, M., Denis, C.S., Richardson, G.P., and Gillespie, P.G. (2004). Hair cells require phosphatidylinositol 4,5-bisphosphate for mechanical transduction and adaptation. Neuron 44:309–320. Howard, J. and Bechstedt, S. (2004). Hypothesis: a helix of ankyrin repeats of the NOMPCTRP ion channel is the gating spring of mechanoreceptors. Current Biology 14:R224–226. Howard, J. and Hudspeth, A.J. (1988). Compliance of the hair bundle associated with gating of mechanoelectrical transduction channels in the bullfrog's saccular hair cell. Neuron 1:189–199. Hudspeth, A.J. (2005). How the ear's works work: mechanoelectrical transduction and ampliﬁcation by hair cells. Comptes Rendus Biologies 328:155–162. Jaquemar, D., Schenker, T., and Trueb, B. (1999). An ankyrin-like protein with transmembrane domains is speciﬁcally lost after oncogenic transformation of human ﬁbroblasts. Journal of Biological Chemistry 274:7325–7333. Jordt, S.E., Bautista, D.M., Chuang, H.H., McKemy, D.D., Zygmunt, P.M., Hogestatt, E.D., Meng, I.D., and Julius, D. (2004). Mustard oils and cannabinoids excite sensory nerve ﬁbres through the TRP channel ANKTM1. Nature 427:260–265. Jorgensen, F. and Ohmori, H. (1988) Amiloride blocks the mechano-electrical transduction channel of hair cells of the chick. Journal of Physiology 403:577–588. Kennedy, H.J., Evans, M.G., Crawford, A.C., and Fettiplace, R. (2003). Fast adaptation of mechanoelectrical transducer channels in mammalian cochlear hair cells. Nature Neuroscience 6:832–836. Kimitsuki, T., Nakagawa, T., Hisashi, K., Komune, S., and Komiyama, S. (1996). Gadolinium blocks mechano-electric transducer current in chick cochlear hair cells. Hearing Research 101:75–80. Kobayashi, K., Fukuoka, T., Obata, K., Yamanaka, H., Dai, Y., Tokunaga, A., and Noguchi, K. (2005). Distinct expression of TRPM8, TRPA1, and TRPV1 mRNAs in rat primary afferent neurons with adelta/c-ﬁbers and colocalization with trk receptors. Journal of Comparative Neurology 493:596–606. Kroese, A.B., Das, A., and Hudspeth, A.J. (1989). Blockage of the transduction channels of hair cells in the bullfrog's sacculus by aminoglycoside antibiotics. Hearing Research 37:203–217. Kumar, G., Duggan, A., and García-Añoveros, J. (2005). TRPA1 channel expression in sensory epithelia of the inner ear and in nociceptors of sensory ganglia. ARO Meeting Abstracts 28:312. Kwan, K.Y., Allchorne, A.J., Vollrath, M.A., Christensen, A.P., Zhang, D.S., Woolf, C.J., and Corey, D.P. (2006). TRPA1 contributes to cold, mechanical, and chemical nociception but is not essential for hair-cell transduction. Neuron 50:277–289. Lee, G., Abdi, K., Jiang, Y., Michaely, P., Bennett, V., and Marszalek, P.E. (2006). Nanospring behaviour of ankyrin repeats. Nature 440:246–249. Lemasurier, M. and Gillespie, P.G. (2005). Hair-cell mechanotransduction and cochlear ampliﬁcation. Neuron 48:403–415. Li, W., Feng, Z., Sternberg, P.W., and Shawn Xu, X.Z. (2006). A C. elegans stretch receptor neuron revealed by a mechanosensitive TRP channel homologue. Nature 440:684–687.

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Macpherson, L.J., Geierstanger, B.H., Viswanath, V., Bandell, M., Eid, S.R., Hwang, S., and Patapoutian, A. (2005). The pungency of garlic: activation of TRPA1 and TRPV1 in response to allicin. Current Biology 15:929–934. Mancillas, J., Duggan, A., and García-Añoveros, J. (2005). Expression and function of TRPA1 channel in Caenorhabditis elegans sensory and non-neuronal cells. SFN Abstracts. Nagata, K., Duggan, A., Kumar, G., and García-Añoveros, J. (2005). Nociceptor and hair cell transducer properties of TRPA1, a channel for pain and hearing. Journal of Neuroscience 25:4052–4061. Obata, K., Katsura, H., Mizushima, T., Yamanaka, H., Kobayashi, K., Dai, Y., Fukuoka, T., Tokunaga, A., Tominaga, M., and Noguchi, K. (2005). TRPA1 induced in sensory neurons contributes to cold hyperalgesia after inﬂammation and nerve injury. Journal of Clinical Investigations 115:2393–2401. Reid, G. (2005). ThermoTRP channels and cold sensing: what are they really up to? Pﬂügers Archives 451:250–263. Ricci, A. (2002). Differences in mechano-transducer channel kinetics underlie tonotopic distribution of fast adaptation in auditory hair cells. Journal of Neurophysiology 87:1738–1748. Ricci, A.J., Crawford, A.C., and Fettiplace, R. (2000). Active hair bundle motion linked to fast transducer adaptation in auditory hair cells. Journal of Neuroscience 20:7131–7142. Ricci, A.J., Crawford, A.C., and Fettiplace, R. (2003). Tonotopic variation in the conductance of the hair cell mechanotransducer channel. Neuron 40:983–990. Rüsch, A., Kros, C.J., and Richardson, G.P. (1994). Block by amiloride and its derivatives of mechano-electrical transduction in outer hair cells of mouse cochlear cultures. Journal of Physiology 474:75–86. Schenker, T. and Trueb, B. (1998). Down-regulated proteins of mesenchymal tumor cells. Experimental Cell Research 239:161–168. Sidi, S., Friedrich, R.W., and Nicolson, T. (2003). NompC TRP channel required for vertebrate sensory hair cell mechanotransduction. Science 301:96–99. Sotomayor, M., Corey, D.P., and Schulten, K. (2005). In search of the hair-cell gating spring elastic properties of ankyrin and cadherin repeats. Structure 13:669–682. Stepanyan, R., Boger, E.T., Friedman, T.B., and Frolenkov, G.L. (2006). TRPA1, a hair cell channel with unknown function? ARO Meeting Abstracts 29:211–212. Story, G.M., Peier, A.M., Reeve, A.J., Eid, S.R., Mosbacher, J., Hricik, T.R., Earley, T.J., Hergarden, A.C., Andersson, D.A., Hwang, S.W., McIntyre, P., Jegla, T., Bevan, S., and Patapoutian, A. (2003). ANKTM1, a TRP-like channel expressed in nociceptive neurons, is activated by cold temperatures. Cell 112:819–829. Tracey, W.D., Jr., Wilson, R.I., Laurent, G., and Benzer, S. (2003). Painless, a Drosophila gene essential for nociception. Cell 113:261–273. Walker, R.G., Willingham, A.T., and Zuker, C.S. (2000). A Drosophila mechanosensory transduction channel. Science 287:2229–2234.

13

TRPM8: The Cold and Menthol Receptor David D. McKemy University of Southern California

CONTENTS Introduction............................................................................................................177 Cold Sensing and Menthol ....................................................................................178 The Hunt for a Menthol Receptor.........................................................................178 The Cloning of a Cold and Menthol Receptor .....................................................180 TRPM8 Has Properties Similar to Those of Cold Receptors ..............................180 Other Cooling Compounds Activate TRPM8 .......................................................182 Regulation of TRPM8 Currents ............................................................................183 Mechanism of Cold Activation..............................................................................184 A Role for TRPM8 Outside the Nervous System ................................................185 Conclusions............................................................................................................186 References..............................................................................................................186

INTRODUCTION Our sensory systems are able to detect subtle changes in ambient temperature, due to the coordinated efforts of thermosensory neurons. At the level of the primary afferent nerve, the site at which thermal stimuli are converted into neuronal activity, temperature-sensitive members of the TRP channel family are found. Remarkably, the range of temperatures that these channels respond to covers the entire perceived temperature spectrum, from warm to painfully hot, from pleasingly cool to excruciatingly cold [1]. Moreover, many of these channels are receptors for ligands that elicit distinct psychophysical sensations, such as the heat associated with capsaicin and the cold felt with menthol. The latter of these was inﬂuential in the discovery of the ﬁrst TRP channel shown to be responsive to temperatures in the cold range (<30°C), TRPM8, a member of the melastatin TRP channel subfamily [2,3]. This chapter focuses on TRPM8, describing what was known about cold signaling before the channel was cloned, how TRPM8 was identiﬁed as a cold sensor, and what advances have been made in our understanding of the molecular logic for cold sensation since its identiﬁcation.

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COLD SENSING AND MENTHOL The perception of nonpainful, cool temperatures is reported to occur when the skin is cooled as little as 1°C from normal body temperature [4]. However, once temperatures approach 15°C, the perception of cold pain is felt, with qualities described as burning, aching, and prickling [5]. In the early to mid-twentieth century, a number of laboratories began to observe cold-induced electrical impulses when recording from mammalian sensory nerves. These peripheral cold receptors, both Aδ- and Cﬁbers, have thermal thresholds (i.e., the temperature at which nerve impulses are generated) for cold activation between 30–20°C, temperatures considered to be innocuously cool [4,6]. Further cooling to temperatures below what is considered noxious (<15°C) was also shown to excite a small percentage of nociceptors (20–30 percent), while cooling to <0°C was reported to activate all ﬁbers [7,8] (for review see reference 9). Thus, there is signiﬁcant diversity in the types of neurons that respond to cold, as well as an expansive range of cold activation thresholds. Moreover, no deﬁned mechanism for cold sensing was described. Most cold-sensitive neurons are also sensitive to the ubiquitous cooling compound menthol, a cyclic terpene alcohol found in mint leaves [10]. It is well known that moderate concentrations of menthol induce a pleasant cool sensation, such as that felt when using menthol-containing products such as candy and vapo-rubs. However, when present at higher doses menthol can be noxious, causing burning, irritation, and pain [10–12]. In seminal studies conducted by Hensel and Zotterman in the 1950s, menthol elicited its “cool” sensation by increasing the threshold temperature for activation of cold receptors [13]. Indeed, the researchers hypothesized that menthol exerted its actions on “an enzyme” that was involved in the activation of these nerves [13]. Surprisingly, it took more than 50 years for Hensel and Zotterman’s hypothesis to be validated.

THE HUNT FOR A MENTHOL RECEPTOR As described previously, studies into cold-sensitive ﬁbers were elusive in deﬁning the biological basis for cold signaling. However, in the mid- to late 1990s, a number of laboratories interested in cold transduction began to use primary cultures of either dorsal root (DRG) or trigeminal (TG) ganglia neurons as in vitro models of sensory afferents. Approximately 10–20 percent of ganglia neurons respond to cold temperatures, with thresholds for activation below 30°C [2,14–16]. Suto and Gotoh showed that cold stimuli (~20°C) evoke a robust inﬂux of calcium in a small percentage (~10 percent) of cultured DRG neurons [15]. Similarly, Kobayashi and colleagues obtained comparable results using menthol as the stimulus, demonstrating that the cooling compound promoted membrane depolarization and generated nerve impulses [17]. Using a similar approach, Reid and Flonta recorded membrane currents from cultured DRG neurons that they selected based on the ability of cold to elicit an increase in cytoplasmic calcium [16]. Cooling generated an inward current in these neurons, when the cells were held at negative membrane potentials, with an average temperature threshold near 29°C. This threshold shifted to warmer temperatures

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B cold

menthone

menthol

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cyclohex

when the recordings were conducted in the presence of menthol, as was predicted from Hensel and Zotterman’s original hypothesis [13]. Similar responses were observed in cultured TG neurons by Julius and colleagues, who went on to demonstrate that both menthol and cold evoke rapidly activating, nonselective cation conductances that were characterized by strong outward rectiﬁcation (Figure 13.1A, B) [2]. Both studies showed that the effects of menthol were temperature dependent and that warming neurons to >37°C could strongly reduce menthol’s effects [2,16].

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FIGURE 13.1 Menthol- and cold-evoked responses in sensory neurons and cells heterologously expressing TRPM8. (A) Whole-cell voltage clamp recordings from dissociated primary cultures of trigeminal neurons. Menthol (100 μM) and cold (see temperature plot) evoke robust membrane currents at both +80 and −60 mV holding potentials. (B) Both menthol(100 μM) and cold-evoked currents display strong outward rectiﬁcation. (C) Recordings from Xenopus oocytes expressing the rat form of TRPM8. Two-electrode voltage-clamp recordings of inward membrane currents (holding potential of −60 mV) evoked when the cells were exposed to 100 μM menthol, 20 mM eucalyptol, and 300 nM AG-3-5. (D) Menthol-(200 μM) and AG-3-5 (2 μM) –evoked currents in TRPM8-expressing HEK293 cells display strong outward rectiﬁcation. Data shown in panels A–D are from McKemy et al [2].

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Both menthol- and cold-evoked currents also adapt to prolonged stimulation at a rate that is similar to what is observed in primates and humans [9]. Thus, this in vitro data supported the hypotheses of Hensel and Zotterman in that it seemed likely that cold and menthol work through a similar mechanism, leading to the search for their common molecular site of action.

THE CLONING OF A COLD AND MENTHOL RECEPTOR The long-sought conﬁrmation of Hensel and Zotterman’s original hypothesis for the action of menthol ﬁnally came in 2002. Two groups working independently and using different experimental approaches concurrently cloned a cold- and mentholsensitive ion channel from sensory neurons [2,3]. The ﬁrst group used menthol to expression clone a complementary DNA (cDNA; a synthesized copy of an RNA transcript) from rat TG neurons that could confer menthol sensitivity to cells that were normally insensitive to temperature [2]. The second group searched for TRP channel–like sequences in mouse DRG neurons and tested these channels for temperature sensitivity [3]. With these divergent approaches, both groups simultaneously identiﬁed TRPM8 (also referred to as trp-p8 or CMR1), a member of the melastatin or long-TRP channel subfamily [18]. Surprisingly, TRPM8 had been identiﬁed prior to these neuronal studies as a transcriptional marker of prostate epithelia, but was not detected in sensory tissue at the time (see below) [19]. Nonetheless, TRPM8 was the ﬁrst cold-activated ion channel to be identiﬁed, and it established the general role for TRP ion channels in thermosensation [1].

TRPM8 HAS PROPERTIES SIMILAR TO THOSE OF COLD RECEPTORS In both TG and DRG, TRPM8 is expressed in <15 percent of small-diameter (~20 μm) sensory neurons, consistent with the proportion of neurons shown to be cold- and menthol-sensitive in neuronal cultures [2,14,20,21]. In initial characterizations of the molecular and neurochemical phenotypes of sensory neurons that express TRPM8, it was found that RNA transcripts for the channel were not coexpressed with the heat-activated TRP channel TRPV1, calcitonin gene-related peptide (CGPR), neuroﬁlament, or isolectin B4 binding [3]. Later studies conﬁrmed these results, further demonstrating that TRPM8 transcripts are in both Aδ- and C-ﬁbers and that it coexpresses with the receptor tyrosine kinase TrkA, but not TrkB or TrkC or another putative temperature-sensitive channel, TRPA1 (see Chapter 11) [22]. Transcripts for TRPM8 are also more abundant in trigeminal versus dorsal root ganglia [2], particularly in the mandibular region that innervates the tongue [22]. TRPM8 was also found to be expressed in lingual nerve ﬁbers that project in to the fungiform papillae of the tongue [23]. Interestingly, TRPM8 ﬁbers were in close proximity to taste buds, but did not innervate these structures. Thus, TRPM8 deﬁnes a small and discrete population of sensory afferents that innervate tissues known to be highly sensitive to cold and nociceptive stimuli.

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TABLE 13.1 Comparison of the Biophysical Properties of Native Cold/Menthol Currents and Heterologously Expressed TRPM8 Native Temperature activation threshold Menthol EC50 Current-voltage relationship Ion permeabilities

Ion selectivity

TRPM8

References

27.1 ± 0.5°; 28.7 ± 2.7°

25.8 ± 0.4°; 21.8 ± 0.6°

80 ± 2.4 μM; Outwardly rectifying

66.7 ± 3.3 μM; 101 ± 13 μM [2]; [30] Outwardly rectifying [2]; [3]

PCa/PNa = 3.2 PK/PNa = 1.1 PCs/PK = 1.2 PCs/PNa = N.A. Erev−Na+ = −5.1 ± 3.1 mV

3.3; 0.97 1.2; 1.34 1.1 1.43 −3.8 ± 2.4 mV; −20 ± 5 mV 0.8 ± 1.0 mV; −5.3 ± 1.5 mV

Erev−K+ = −4.5 ± 2.0 mV

Erev−NMDG = −85.0 ± 11.5 mV Erev−Ca2+−NMDG = −43.3 ± 4.6 mV −77.2 ± 5.9 mV −35.2 ± 8.0 mV

[2]; [16]; [3]

[2]; [3] [2]; [3] [2] [3] [2]; [48] [2]; [48] [2] [2]

When expressed in heterologous expression systems, such as Xenopus oocytes or mammalian cell lines, TRPM8-mediated currents are activated by a number of cooling compounds in addition to menthol, such as eucalyptol (the active ingredient in eucalyptus oil) and the super-cooling AG-3-5 (Figure 13.1C) [2]. Biophysically, TRPM8 has surprisingly similar properties to those recorded in both cultured DRG and TG using similar experimental paradigms (see Table 13.1). These include selectivity for ions, potency of menthol in activating currents, and voltage dependence of membrane currents induced by either cold or menthol [2,3,16,24]. Like almost all TRP channels, TRPM8 is a nonselective cation channel that displays strong outward rectiﬁcation (Figure 13.1D). Like native menthol-evoked currents, TRPM8 showed relatively high selectivity for calcium and little selectivity among monovalent cations [2]. Menthol-evoked single-channel currents are also characterized by strong outward rectiﬁcation and have a slope conductance of 83 pS [2]. More remarkably, TRPM8 currents are also evoked by temperature decreases with an activation temperature threshold of ~26°C, with activity increasing in magnitude down to 8°C (Figure 13.2A, B) [2,3]. Interestingly, this broad range spans what are considered both innocuous cool (~30–15°C) and noxious cold temperatures (<15°C). Moreover, neither the rate nor direction of temperature change altered the response proﬁle of TRPM8 currents (Figure 13.2B) [2]. Therefore, the channel’s response is directly proportional to the temperature presented in these restricted in vitro systems. When menthol is applied, the threshold for activation shifts to warmer temperatures (Figure 13.2C) [2], and increasing

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Temperature (oC) Temperature (oC) FIGURE 13.2 Cold activation of TRPM8. (A) Reduction in perfusate temperature (from ~32 to 10°C) evokes robust inward currents in TRPM8-expressing Xenopus oocytes (−60 mV hp.). Two rates of temperature change are shown (0.2°C/sec.; 1°C/sec.). (B) Temperature-response proﬁle of the currents shown in A. (C) Independent recordings from several TRPM8-expressing oocytes show an activation temperature of ~26°C. Temperature ramps performed in the presence of 20 μM menthol shift the threshold to warmer temperatures. (D) Increasing concentrations of menthol shift the temperature response proﬁle to warmer temperatures. Moreover, even at high menthol concentrations, TRPM8 currents are inhibited by warm temperatures. Data shown in panels A–C are from McKemy et al. [2], and data in panel D are unpublished observations (D. McKemy).

concentrations of menthol shift this curve even more, suggesting that menthol mimics the endogenous mechanism for thermal activation of TRPM8 (Figure 13.2D; D. McKemy, unpublished observations).

OTHER COOLING COMPOUNDS ACTIVATE TRPM8 In addition to menthol and AG-3-5, a number of cooling agents, including CoolactP, Cooling Agent 10, FrescolatMGA, FrescolatML, geraniol, hydroxycitronellal, linalool, PMD38, WS-3, and WS-23 activate TRPM8 in vitro [2,25,26]. Of these

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AG-3-5 (also known as icilin) was ﬁrst identiﬁed as a super-cooling agent in the early 1980s and bears little resemblance to menthol structurally [27]. AG-3-5 is more potent and effective than menthol in activating TRPM8, and when given intravenously, it will induce characteristic shivering or “wet dog” shakes [2,27]. Interestingly, the mechanism whereby AG-3-5 activates TRPM8 is different than that of menthol or cold [2]. AG-3-5 requires a coincident rise in cytoplasmic calcium, either via permeation through the channel or by release from intracellular stores, in order to evoke TRPM8 currents [28]. This requirement of a calcium rise for TRPM8 activity is not needed for cold- or menthol-induced channel activity, suggesting the channel can be activated by multiple mechanisms. Additionally, a critical amino acid was identiﬁed: when mutated, it rendered AG-3-5 incapable of activating TRPM8. This residue was located between the second and third transmembrane domains of the channel, a region known to be important for capsaicin sensitivity of TRPV1 [28,29]. While various compounds activate TRPM8, a more relevant class of molecules that may be of use clinically is of those that antagonize or block the channel. A number of antagonists have been identiﬁed, including BCTC, thio-BCTC, CTPC, and capsazepine [26,30]. Surprisingly, many of these compounds also antagonize the heat-gated channel TRPV1. Thus, there is signiﬁcant overlap pharmacologically between the two channels. Along with the analogous positions critical for capsaicin and AG-3-5 activation of TRPV1 and TRPM8, respectively, these results suggest a conserved mechanism for ligand activation of these thermosensitive TRP channels. While of interest pharmacologically, these results complicate the search for selective agents for these channels that are such good targets for drug discovery.

REGULATION OF TRPM8 CURRENTS Experience tells us that temperature sensation is a dynamic process. For instance, we can easily adapt to cold temperatures, a process observed in both psychophysical and cellular assays [6,16,24,31]. Similarly, cold- or menthol-induced TRPM8 currents will adapt or desensitize in a calcium-dependent manner during prolonged stimulation [2]. Increased intracellular calcium is known to lead to the breakdown of a membrane phospholipid, phosphatidylinositol 4,5-bisphosphate (PIP2), via activation of phospholipase C (PLC) [32]. Thus, it has been proposed that calcium inﬂux, via TRPM8, activates PLC to cleave PIP2, thereby altering the concentration of this phospholipid in the plasma membrane [33,34]. Thus, adaptation is thought to be due to cellular changes in the levels of PIP2, which modulates the ability of TRPM8 to respond to cold or menthol. At the molecular level, a number of amino acid residues have been identiﬁed in the carboxy-terminal domain of the channel, adjacent to the sixth transmembrane domain, that appear to be involved in PIP2’s effects on TRPM8 [34]. Interestingly, these residues are near the highly conserved TRP box of the channel and are found in other PIP2-sensitive TRPM channels, including TRPM4 and TRPM5 [35,36]. In addition to calcium, many types of pathological conditions, such as peripheral inﬂammation, can lead to altered membrane levels of PIP2 [32]. Many cell-surface receptors, such as TrkA, the receptor for nerve growth factor, and the bradykinin

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receptor activate PLC, thereby cleaving PIP2 in the membrane. A consequence of PIP2 breakdown is the generation of diacylglycerol (DAG) and inositol-trisphosphate (IP3). DAG, along with increased intracellular calcium, activates protein kinase C (PKC), leading to phosphorylation of several cellular substrates including many ion channels. Phosphorylation is a common mechanism whereby channel activity is modulated, and increased PKC activity causes decreased TRPM8 membrane currents [37]. Interestingly, PKC activation does not lead to increased incorporation of phosphate on TRPM8, but rather a decrease in TRPM8 phosphorylation. This was blocked by treatment with phosphatase inhibitors, suggesting that the PKC-mediated effects are not due to direct phosphorylation of TRPM8, but that PKC plays a role upstream of channel phosphorylation. Intracellular pH also regulates TRPM8. When pH is increased to above physiological levels, TRPM8 activity is inhibited [26,30]. These effects of pH are thought to be mediated intracellularly [30], but there is disagreement on the effects of pH on cold-, menthol-, and AG-3-5–evoked currents. Andersson et al. reported that menthol’s ability to activate TRPM8 is unaffected by pH, but that cold and AG-3-5 responses are inhibited [30]. Behrendt et al. also found that AG-3-5 was less effective in activating TRPM8 at high pH, but in contrast, menthol-evoked responses were also suppressed [26]. Cold was not tested in the latter study. Thus, it seems likely that either cell-to-cell variation in temperature thresholds for cold, or altered sensitivity due to experience and pathological state of the neuron, may be a result of TRPM8 regulation via cellular levels of PIP2, protons, or kinase activity. This level of channel modulation may also account for the complexity and variability in cold-evoked temperature responses observed both in vivo and in vitro.

MECHANISM OF COLD ACTIVATION Ever since the identiﬁcation of thermosensitive TRP ion channels, the basis of the precipitous temperature sensitivity of these proteins has been of keen interest, but a physiological or molecular mechanism has remained elusive. Several plausible mechanisms have been proposed, including temperature-dependent structural reorganization of the channels, production of endogenous channel-activating ligands by a change in temperature, or that the channels respond to temperature-dependent changes in membrane ﬂuidity [38]. For TRPM8, temperature has been shown to produce two fundamental changes in channel properties: a shift in the voltage dependence of the channel and modiﬁcation of the maximum probability of channel opening [39,40]. Like other members of the TRPM subfamily, TRPM8 currents are voltage sensitive [2], and an initial study reported that declining temperatures shift the voltage-dependent activation curve of TRPM8 toward negative membrane potentials [40]. Kinetic analyses suggested that temperature sensitivity of the channel was a result of an over ten-fold difference between the activation energies associated with opening and closing the channel (Ea,open < Ea,close). Thus, cold activation of TRPM8 was described with a simple, single-thermodynamic principle based on this difference and was not due to

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signiﬁcant changes in the steady-state open probability of the channel. Moreover, menthol was shown to mimic the effect of cold temperatures on voltage dependence of the channel, serving as a gating modiﬁer [40]. A subsequent report also provided evidence for a temperature-dependent shift in the voltage dependence of TRPM8 [39]. However, in contrast to the previous ﬁndings, a signiﬁcant increase in the maximum open probability of the channel was observed upon cooling, leading to the hypothesis that temperature affects not only the voltage dependence of the channel, but also leads to large conformational changes in TRPM8. Thus, a more complex allosteric model for channel activation was proposed, one that suggests that TRPM8 has two autonomous sensors, one for voltage and one for temperature, that are activated independently and interact to promote channel opening [39]. To date, the structural determinants of voltage and temperature sensitivity of TRPM8 have not been identiﬁed. However, the similarity of the TRP channel family to classic voltage-gated channels suggests that a putative voltage sensor may reside in the fourth transmembrane domain [41]. Whether the temperature sensor exists in this or other regions of the channel remains to be determined.

A ROLE FOR TRPM8 OUTSIDE THE NERVOUS SYSTEM Although TRPM8 is undoubtedly critical for transduction of thermal stimuli in the peripheral nervous system, its expression in other tissues suggests it serves other biological roles in addition to neuronal thermal sensing. Along with expression in the prostate, TRPM8 has also been found in the bladder and male genital tract [42]. It is also observed that TRPM8 expression increases dramatically in cancers of the prostate, as well as other nonprostatic tumors such as breast, colon, lung, and skin [19]. In the prostate, low levels of TRPM8 transcripts have been identiﬁed [19]. However, TRPM8 expression greatly increases in transformed prostate epithelia. A role for TRPM8 in either normal or cancerous prostate tissues is still unclear, and studies using models of prostate cancer, such as LNCaP cells, have shown that the channel does respond to cold and menthol in these cells [43]. However, the channel is not present on the cell surface but is trapped in intracellular membrane compartments such as the endoplasmic reticulum. This expression is androgen dependent, suggesting that TRPM8 may play a role in differentiating these cells [44,45]. In addition to prostate and cancerous tissues and cells, TRPM8 transcripts have been found in the gastric fundus [46]. Cooling that takes place following the consumption of cold foods induces contraction of gastrointestinal smooth muscles, resulting in a short-lived gastric voiding [47]. When a putative TRPM8 inhibitor capsazepine was applied, these cooling-evoked contractions were reduced, suggesting that TRPM8 may be involved in this process in a nonneuronal capacity [46]. However, capsazepine is also a potent antagonist for TRPV1, thus complicating these results. Nonetheless, these and other data strongly suggest that TRPM8 may have diverse biological functions outside of the peripheral nervous system.

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CONCLUSIONS The elucidation of TRP channels as molecular detectors of thermal stimuli addressed a fundamental issue in sensory transduction: how are thermal stimuli converted into neuronal activity? The identiﬁcation of TRPM8 and the subsequent studies characterizing its properties have shed light on the molecular mechanisms of cold sensation. Future studies will undoubtedly continue to elucidate the biological importance of this ion channel in mediating sensory signaling, as well as the role of TRPM8 in nonneuronal tissues. Nonetheless, cloning TRPM8 established the ﬁrst molecular detector of cold stimuli, and its in vitro properties are consistent with this role in vivo. Moreover, TRPM8 conﬁrmed Hensel and Zotterman’s half-century-old hypothesis [13] and established that TRP channels can confer thermal stimuli over broad ranges of temperature.

REFERENCES 1. Jordt, S.E., D.D. McKemy, and D. Julius, Lessons from peppers and peppermint: the molecular logic of thermosensation. Current Opinion in Neurobiology, 2003(13). 2. McKemy, D.D., W.M. Neuhausser, and D. Julius, Identiﬁcation of a cold receptor reveals a general role for TRP channels in thermosensation. Nature, 2002. 416(6876): 52–58. 3. Peier, A.M. et al., A TRP channel that senses cold stimuli and menthol. Cell, 2002. 108(5): 705–15. 4. Campero, M. et al., Slowly conducting afferents activated by innocuous low temperature in human skin. J. Physiol., 2001. 535(Pt 3): 855–65. 5. Morin, C. and M.C. Bushnell, Temporal and qualitative properties of cold pain and heat pain: a psychophysical study. Pain, 1998. 74(1): 67–73. 6. Darian-Smith, I., K.O. Johnson, and R. Dykes, ‘‘Cold’’ ﬁber population innervating palmar and digital skin of the monkey: responses to cooling pulses. J. Neurophysiol., 1973. 36(2): 325–46. 7. Simone, D.A. and K.C. Kajander, Excitation of rat cutaneous nociceptors by noxious cold. Neurosci. Lett., 1996. 213(1): 53–56. 8. Simone, D.A. and K.C. Kajander, Responses of cutaneous A-ﬁber nociceptors to noxious cold. J. Neurophysiol., 1997. 77(4): 2049–60. 9. Reid, G., ThermoTRP channels and cold sensing: what are they really up to? Pﬂügers Arch., 2005. 10. Eccles, R., Menthol and related cooling compounds. J. Pharm. Pharmacol., 1994. 46(8): 618–30. 11. Cliff, M.A. and B.G. Green, Sensory irritation and coolness produced by menthol: evidence for selective desensitization of irritation. Physiol. Behav., 1994. 56(5): 1021–29. 12. Green, B.G., The sensory effects of l-menthol on human skin. Somatosens. Mot. Res., 1992. 9(3): 235–44. 13. Hensel, H. and Y. Zotterman, The effect of menthol on the thermoreceptors. Acta Physiol. Scand., 1951. 24: 27–34. 14. Viana, F., E. de la Pena, and C. Belmonte, Speciﬁcity of cold thermotransduction is determined by differential ionic channel expression. Nat. Neurosci., 2002. 5(3): 254–60.

TRPM8: The Cold and Menthol Receptor

187

15. Suto, K. and H. Gotoh, Calcium signaling in cold cells studied in cultured dorsal root ganglion neurons. Neuroscience, 1999. 92(3): 1131–35. 16. Reid, G. and M.L. Flonta, Physiology. Cold current in thermoreceptive neurons. Nature, 2001. 413(6855): 480. 17. Okazawa, M. et al., l-Menthol-induced [Ca2+]i increase and impulses in cultured sensory neurons. Neuroreport, 2000. 11(10): 2151–55. 18. Moran, M.M., H. Xu, and D.E. Clapham, TRP ion channels in the nervous system. Curr. Opin. Neurobiol., 2004. 14(3): 362–69. 19. Tsavaler, L. et al., Trp-p8, a novel prostate-speciﬁc gene, is up-regulated in prostate cancer and other malignancies and shares high homology with transient receptor potential calcium channel proteins. Cancer Res., 2001. 61(9): 3760–69. 20. Reid, G. and M.L. Flonta, Ion channels activated by cold and menthol in cultured rat dorsal root ganglion neurones. Neurosci. Lett., 2002. 324(2): 164–68. 21. Thut, P.D., D. Wrigley, and M.S. Gold, Cold transduction in rat trigeminal ganglia neurons in vitro. Neuroscience, 2003. 119(4): 1071–83. 22. Kobayashi, K. et al., Distinct expression of TRPM8, TRPA1, and TRPV1 mRNAs in rat primary afferent neurons with adelta/c-ﬁbers and colocalization with trk receptors. J. Comp. Neurol., 2005. 493(4): 596–606. 23. Abe, J. et al., TRPM8 protein localization in trigeminal ganglion and taste papillae. Brain Res. Mol. Brain Res., 2005. 136(1–2): 91–98. 24. Reid, G., A. Babes, and F. Pluteanu, A cold- and menthol-activated current in rat dorsal root ganglion neurons: properties and role in cold transduction. J. Physiol., 2002. 545(Pt. 2): 595–614. 25. Weil, A. et al., Conservation of functional and pharmacological properties in the distantly related temperature sensors TRVP1 and TRPM8. Mol. Pharmacol., 2005. 68(2): 518–27. 26. Behrendt, H.J. et al., Characterization of the mouse cold-menthol receptor TRPM8 and vanilloid receptor type-1 VR1 using a ﬂuorometric imaging plate reader (FLIPR) assay. Br. J. Pharmacol., 2004. 141(4): 737–45. 27. Wei, E.T. and D.A. Seid, AG-3-5: a chemical producing sensations of cold. J. Pharm. Pharmacol., 1983. 35(2): 110–12. 28. Chuang, H.H., W.M. Neuhausser, and D. Julius, The super-cooling agent icilin reveals a mechanism of coincidence detection by a temperature-sensitive TRP channel. Neuron, 2004. 43(6): 859–69. 29. Jordt, S.E. and D. Julius, Molecular basis for species-speciﬁc sensitivity to ‘‘hot’’ chili peppers. Cell, 2002. 108(3): 421–30. 30. Andersson, D.A., H.W. Chase, and S. Bevan, TRPM8 activation by menthol, icilin, and cold is differentially modulated by intracellular pH. J. Neurosci., 2004. 24(23): 5364–69. 31. Kenshalo, D.R. and R. Duclaux, Response characteristics of cutaneous cold receptors in the monkey. J. Neurophysiol., 1977. 40(2): 319–32. 32. Hilgemann, D.W., S. Feng, and C. Nasuhoglu, The complex and intriguing lives of PIP2 with ion channels and transporters. Sci. STKE, 2001. 2001(111): RE19. 33. Liu, B. and F. Qin, Functional control of cold- and menthol-sensitive TRPM8 ion channels by phosphatidylinositol 4,5-bisphosphate. J. Neurosci., 2005. 25(7): 1674–81. 34. Rohacs, T. et al., PI(4,5)P2 regulates the activation and desensitization of TRPM8 channels through the TRP domain. Nat. Neurosci., 2005. 8(5): 626–34. 35. Zhang, Z. et al., Phosphatidylinositol 4,5-bisphosphate rescues TRPM4 channels from desensitization. J. Biol. Chem., 2005. 280(47): 39185–92.

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36. Liu, D. and E.R. Liman, Intracellular Ca2+ and the phospholipid PIP2 regulate the taste transduction ion channel TRPM5. Proc. Natl. Acad. Sci. USA, 2003. 100(25): 15160–65. 37. Premkumar, L.S. et al., Downregulation of transient receptor potential melastatin 8 by protein kinase C–mediated dephosphorylation. J. Neurosci., 2005. 25(49): 11322–29. 38. Clapham, D.E., TRP channels as cellular sensors. Nature, 2003. 426(6966): 517–24. 39. Brauchi, S., P. Orio, and R. Latorre, Clues to understanding cold sensation: thermodynamics and electrophysiological analysis of the cold receptor TRPM8. Proc. Natl. Acad. Sci. USA, 2004. 101(43): 15494–99. 40. Voets, T. et al., The principle of temperature-dependent gating in cold- and heatsensitive TRP channels. Nature, 2004. 430(7001): 748–54. 41. Hille, B., Ion channels of excitable membranes. 3rd ed. 2001, Sunderland, Mass.: Sinauer. 42. Stein, R.J. et al., Cool (TRPM8) and hot (TRPV1) receptors in the bladder and male genital tract. J. Urol., 2004. 172(3): 1175–78. 43. Thebault, S. et al., Novel role of cold/menthol-sensitive TRPM8 in the activation of store-operated channels in LNCaP human prostate cancer epithelial cells. J. Biol. Chem., 2005. 44. Bidaux, G. et al., Evidence for speciﬁc TRPM8 expression in human prostate secretory epithelial cells: functional androgen receptor requirement. Endocr. Relat. Cancer, 2005. 12(2): 367–82. 45. Zhang, L. and G.J. Barritt, Evidence that TRPM8 is an androgen-dependent Ca2+ channel required for the survival of prostate cancer cells. Cancer Res., 2004. 64(22): 8365–73. 46. Mustafa, S. and M. Oriowo, Cooling-induced contraction of the rat gastric fundus: mediation via transient receptor potential (TRP) cation channel TRPM8 receptor and Rho-kinase activation. Clin. Exp. Pharmacol. Physiol., 2005. 32(10): 832–38. 47. Mustafa, S.M. and O. Thulesius, Cooling-induced gastrointestinal smooth muscle contractions in the rat. Fundam. Clin. Pharmacol., 2001. 15(5): 349–54. 48. Hui, K., Y. Guo, and Z.P. Feng, Biophysical properties of menthol-activated cold receptor TRPM8 channels. Biochem. Biophys. Res. Commun., 2005. 333(2): 374–82.

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Activation Mechanisms and Functional Roles of TRPP2 Cation Channels Aurélie Giamarchi and Patrick Delmas Laboratoire de Neurophysiologie Cellulaire, CNRS

CONTENTS Abstract ..................................................................................................................189 Introducing the TRPP Protein Subfamily .............................................................190 Mutations in TRPP2 and Polycystin-1 Cause ADPKD ........................................191 Localization and Trafﬁcking of TRPP2 ................................................................193 TRPP2: A Ca2+-Regulated Cation Channel Located in the Endoplasmic Reticulum ..............................................................................194 Coassembly of PKD1 and TRPP2 Reconstitutes a Cell Surface Ca2+-Permeable Cation Channel with Multiple Functions ...................................194 Activation of the PKD1–TRPP2 Complex ...........................................................195 TRPP2: A Bona Fide Mechanosensitive Channel?...............................................196 Concluding Remarks .............................................................................................199 Acknowledgments..................................................................................................199 References..............................................................................................................199

ABSTRACT TRPP2 (polycystin-2) is a member of the TRP family of nonselective cation channels that is mutated in human autosomal polycystic kidney disease. TRPP2 has been implicated in Ca2+-dependent mechanosensitive pathways in a variety of biological functions including cell proliferation, sperm fertilization, mating behavior, and asymmetric gene expression. Although its function as a Ca2+-permeable cation channel is well established, its precise role and subcellular localization in the plasma membrane, endoplasmic reticulum (ER), and cilium have remained controversial. The present review summarizes the most pertinent recent evidence regarding the structural and functional properties of TRPP2 channels, focusing on the regulation and physiology of mammalian TRPP2.

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INTRODUCING THE TRPP PROTEIN SUBFAMILY The transient receptor potential (TRP) channel superfamily currently includes 56 related six-transmembrane (TM) domain channels classiﬁed in seven subfamilies designated TRPC (“canonical”), TRPV (“vanilloid”), TRPM (“melastatin”), TRPN (“NOMPC,” from no mechanoreceptor potential-C), TRPA (“ankyrin-like with transmembrane domain-1”), TRPML (“mucolipin”), and TRPP (“polycystin”) [1,2]. TRPC, TRPV, and TRPM are related to canonical TRP proteins while TRPN, TRPA, and TRPP are more divergent. TRP channels are linked to a variety of sensory stimuli including phototransduction, thermosensation, and mechanosensation and to multiple integrative cellular functions including Ca2+ and Mg2+ homeostasis and cell cycle. The TRPP subfamily was named after its founding member polycystin kidney disease-2 (TRPP2 encoded by the PKD2 gene), a gene product mutated in an inherited human disorder known as autosomal dominant polycystic kidney disease (ADPKD). TRPP2 is related to the TRP family of ion channels by virtue of its topological features/structural homology in the sixth TM region and the presence of an ion channel motif. The polycystin proteins are found in the entire animal kingdom. In humans, the polycystin group contains eight members, which are widely expressed and can be divided structurally into two prototypical subgroups: the PKD1-like proteins and the PKD2 (TRPP2)-like proteins, both having a modest degree of sequence similarity in their C-termini [3–6]. The TRPP subfamily contains three homologous proteins, PKD2, PKD2L1, and PKD2L2, which are currently referred to as TRPP2, TRPP3, and TRPP5 [7]. The mammalian orthologues are highly conserved over the entire length (80–90 percent identity) but show sequence divergence in their cytosolic N-terminal regions. All TRPP2-like proteins are predicted to possess a putative coiled-coil domain at their C-termini, but only TRPP2 and TRPP3 have a Ca2+-binding EF-hand motif. The TRPP2 C-terminus also has a PACS-interacting binding motif, which plays a key role in regulating TRPP2 trafﬁcking between organelles and the cell surface (reference 9 and see below). TRPP3 was the ﬁrst PKD2-like protein to be identiﬁed as a functional TRP-like cation channel [8]. TRPP2-related channels have large single-channel conductance (80–160 pS) and permeate a number of mono- and divalent cations, including Na+, K+, Ba2+, and Ca2+ [reviewed in reference 10]. Like many TRP channels, TRPP2 activity is blocked by La3+ and reduced by the diuretic amiloride [10]. TRPP3 is usually found to be more permeable to Ca2+ than to other monovalent cations such as Na+ and K+ (PCa/PNa = 4), whereas TRPP2 has a PCa /PNa selectivity ranging from 1 to 3. TRPP2-like cation channels therefore represent relevant routes for calcium entry or release. PKD1, PKD1L13, and PKDREJ (receptor for egg jelly) are large multidomain proteins and make up the PKD1-like subgroup. Because PKD1-like proteins show very limited sequence similarity with TRP channels, they are not considered members of the TRP superfamily. Each PKD1-like protein contains a large extracellular terminal domain, 11 predicted TM segments, and a short intracellular

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carboxyl terminus. They all possess the combination of REJ (receptor for egg jelly), GPS (G-protein-coupled receptor proteolytic site), and PLAT/LH2 (lipoxygenase homology/polycystin, lipoxygenase, α-toxin) domains that uniquely deﬁne them as PKD1 family members. The extracellular region of PKD1 spans more than ~3,000 amino acids and contains a number of adhesive domains that implicate PKD1 in cell–cell and cell–matrix interactions. PKD1 is cleaved at its predicted GPS [11], a feature common to members of the family-B (latrophilin) G-proteincoupled receptors and which may be important for receptor activation or localization. PKD1, PKD1L1, and PKD1L2 encompass a G-protein-interacting site that may be used to regulate at least four different classes of heterotrimeric Gprotein activity and multiple downstream effectors, including among others phospholipase C, protein kinase C, adenylyl cyclase, protein kinase A, Janus kinase 2, and nuclear factor of activated T cells (NFAT) [reviewed in reference 4] (Figure 14.1). The intracellular carboxyl terminus of PKD1 also harbors a coiledcoil domain that is involved in physical interaction with TRPP2 [12] and possibly TRPP3a [13]. PKD1 and PKD1L1, but not PKDREJ, PKD1L2, or PKD1L3, have this coiledcoil domain, suggesting that the PKD1-like family could be subdivided into two groups according to their structural domains. Likewise, though PKD1 and PKD1L1 lack a structurally deﬁned surface channel pore domain, recent data suggest that PKD1L2, PKD1L3, and PKDREJ contain strong ion channel signature motifs [14], suggesting their potential functions as pore-forming channel subunits.

MUTATIONS IN TRPP2 AND POLYCYSTIN-1 CAUSE ADPKD The founding members of the TRPP family were discovered as genes mutated in ADPKD. ADPKD is a common nephropathy affecting 4 to 6 million people worldwide and is a leading cause of end-stage kidney failure. The disease is typically characterized by defects in the polarized phenotype and function of epithelial kidney cells, leading to abnormal renal tubular cell growth and formation of numerous ﬂuidﬁlled cysts. Eventually these cysts overwhelm the kidney and destroy the parenchyma. This disease is as yet incurable and is often associated with a number of systemic manifestations including hypertension, intracranial aneurysms, and cardiovalvular abnormalities such as mitral valve prolapse. In the mid-1990s, mutated genes responsible for ADPKD were identiﬁed by positional cloning [15]. ADPKD has been shown to result from loss-of-function mutations either of polycystin-1 or of TRPP2, with PKD1 mutations being the most prevalent causes. The dominance of PKD1 and TRPP2 mutations appears to require both a germ-line mutation of PKD1 or TRPP2 and a subsequent somatic mutation of the wild-type allele. This would explain the relatively late development of ADPKD and the focal nature of epithelial cells giving rise to cysts. However, recent studies indicate that various karyotypic changes, not just loss of heterozygosity at the normal

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FIGURE 14.1 (Color ﬁgure follows p. 234.) Signaling and regulation of TRPP2 channels. This illustration depicts the different models for the localization-dependent functions of TRPP2. TRPP2 mediates Ca2+ inﬂux at the plasma membrane (PM) and the ciliary membrane (cilium), where it functions in a heteromultimeric protein complex with PKD1 (1, 3) and possibly with other members of the TRP channel superfamily (TRPC1, TRPV4) (2). PKD1 is known to activate multiple signaling pathways via G proteins, a process that can be regulated by physical interaction with TRPP2. Calcium inﬂux induced by shear ﬂow in renal epithelial cells is triggered by bending the primary cilium and seemingly requires the PKD1–TRPP2 complex in the luminal cilium (3). Mechanical stress activates PKD1–TRPP2 complexes to allow Ca2+ inﬂux either in the shaft or in the base of the primary cilium. TRPP2 acts as a Ca2+-release channel in the endoplasmic reticulum (ER), where it might interact with and regulate IP3Rs (4). Serine-phosphorylated TRPP2 sequesters Id2 in the cytoplasm and normally prevents Id2 from entering the nucleus and binding to E-proteins (5). PKD1 can undergo a proteolytic cleavage that releases its C-terminal tail, which translocates to the nucleus and activates the transcription factor AP-1 (6). Note that we hypothetized that PKD1 targeted to the primary cilium is also cleaved at its GPS site. See text for more details. Abbreviations: PKC, protein kinase C; PKA, cAMP-dependent protein kinase; NFAT, nuclear factor of activated T-cells; PI3K, phosphatidyl-inositol 3-kinase; MEK, mitogen-activated protein kinase/ERK kinase; ERK, extracellular signal-regulated kinase.

PKD allele, are associated with cystogenesis [16], a situation that illustrates the complexity of cyst formation and raises the question as to whether the two-hit mechanism is the only means to generate a cyst. Consistent with the broad expression of both genes during early organogenesis, mouse models for ADPKD derived from targeted disruption of either PKD genes

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die in utero or perinatally with cardiac septal defects and severe cystic manifestations in nephrons and pancreatic ducts.

LOCALIZATION AND TRAFFICKING OF TRPP2 TRPP2 is expressed in a variety of tissues including epithelial cells, vascular smooth muscle, cardiac myocytes, adrenal glands, and ovaries [17]. A long-lasting matter of debate has been the subcellular localization of TRPP2. In recombinant cell-based systems and in most native cells, TRPP2 is found to be concentrated in intracellular compartments and most notably in the ER, as judged by immunoﬂuorescence imaging, cofractionation with ER markers, and sensitivity to endoglycosidase H digestion [18]. TRPP2 encompasses an ER retention signal within its C-terminal domain [18], which seems to prevent trafﬁcking to the cell surface when expressed on its own. Deletion mutants for this ER retention signal translocate to the cell surface and can be detected by immunological and electrophysiological means [19]. Opposing these ﬁndings are reports that electrogenic TRPP2 activity is present at the plasma membrane of several cell types following treatment with chemical chaperones/proteasome inhibitors [20,21] or upon overexpression [22]. TRPP2 has also been localized to basolateral plasma membranes, lamelopodia, primary cilia, and mitotic spindles [12,18,23–26]. Recent data are clarifying the confusing picture of TRPP2 localization. First, Köttgen et al. [9] have reported that TRPP2 trafﬁcking between the ER, Golgi, and plasma membrane compartments may be directed by the phosphofurin acidic cluster proteins PACS-1 and PACS-2, two sorting proteins that bind to an acidic cluster in the C-ter domain of TRPP2. Binding of these adaptor proteins to TRPP2 is promoted by protein kinase casein kinase-2-dependent phosphorylation of Ser812. TRPP2 accumulates at the plasma membrane only when both PACS-1 and PACS-2 molecules are absent or upon inhibition of protein kinase casein kinase-2 activity. Mutation of Ser812 to alanine or destruction of the acidic cluster (TRPP2 D815–817A) abrogates the interaction between TRPP2 and PACS proteins and increases whole-cell TRPP2 currents. Thus, mechanisms that regulate the interaction of PACS proteins with TRPP2 are likely to play key roles in routing TRPP2 between the ER and the plasma membrane. TRPP3 and TRPP5 lack the PACSbinding acidic cluster in their C-tails, suggesting that their trafﬁcking is regulated differently from that of TRPP2. This may explain why TRPP3 is targeted to the cell surface when overexpressed in oocytes, while TRPP2 is retained in the ER. Second, a recently discovered protein of 14 kDa dubbed PIGEA-14 (polycystin-2 interactor, Golgi- and ER-associated protein) has been shown to interact with the C-ter of TRPP2. Coexpression of both proteins in HeLa cells and in LLC-PK1 induces a redistribution of TRPP2 as well as PIGEA-14 from the ER to an unorthodox trans-Golgi compartment [27], indicating that intracellular trafﬁcking of TRPP2 is regulated both at the levels of the ER and the trans-Golgi network. Trafﬁcking of TRPP2 is therefore rapidly becoming a key issue to understanding TRPP2 function and dysfunction [28].

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TRPP2: A CA2+-REGULATED CATION CHANNEL LOCATED IN THE ENDOPLASMIC RETICULUM TRPP2 may act as a Ca2+-release channel in ER membranes, which ampliﬁes Ca2+ transients initiated by InsP3-generating plasma membrane receptors (Figure 14.1) [29]. This led to the suggestion that TRPP2 represents a new type of intracellular receptor that, along with IP3Rs and RyRs, may be involved in mediating Ca2+-induced Ca2+ release. TRPP2 appears to be directly activated by Ca2+ and displays a bellshaped dependence on cytoplasmic Ca2+ [29, see 10 for review]. Although it is yet unclear whether the Ca2+-binding EF hand of TRPP2 is involved in Ca2+-dependent modulation of TRPP2, it is noteworthy that pathogenic TRPP2 mutants with premature termination of the peptide chain in their C-ter lost their ability to sense Ca2+. However, TRPP3 C-terminal artiﬁcial truncation mutants lacking the EF hand exhibit basal and Ca2+-activated channel activities, suggesting that, at least for TRPP3, the EF hand and other parts of the carboxyl tail are not key determinants of the Ca2+-dependent activation [30]. Phosphorylation of Ser812 by a putative casein kinase 2 results in a signiﬁcant increase in the sensitivity of the TRPP2 channel to calcium stimulation [31]. The S812A substitution, which results in loss of phosphorylation of TRPP2, shifts the Ca2+ dependence such that TRPP2 S812A has a maximum open probability at tenfold higher Ca2+ concentrations (∼3 μM [Ca2+]) than normal TRPP2. Thus, the maximum open probability of a wild-type TRPP2 channel occurs at a Ca2+ concentration at which the nonphosphorylated form remains closed. Intracellular TRPP2 is therefore likely to have enhanced Ca2+ sensitivity, because the protein kinase casein kinase-2 is opportunely associated with the ER and most TRPP2 is found to be phosphorylated in vivo [31]. As stated above, phosphorylated TRPP2 fails to escape the ER. Only dephosphorylation of S812 would promote TRPP2 translocation to the plasma membrane. Thus, it can be hypothesized that the dephosphorylated form of TRPP2, possibly the form found in the plasma or the ciliary membranes, is unlikely to be activated by a modest increase in bulk Ca2+. In line with a role of ER-localized TRPP2 in regulating intracellular Ca2+, TRPP2+/- vascular smooth muscle cells show a lower level of TRPP2 and have altered intracellular Ca2+ homeostasis [32]. Furthermore, TRPP2 has been recently shown to interact functionally and physically with IP3R in oocyte expression systems (Figure 14.1) [33]. The physiological relevance of these results remains to be clariﬁed.

COASSEMBLY OF PKD1 AND TRPP2 RECONSTITUTES A CELL SURFACE CA2+-PERMEABLE CATION CHANNEL WITH MULTIPLE FUNCTIONS The differential cellular and subcellular pattern of expression of PKD1 and TRPP2 in some systems strongly argues that both proteins have independent functions [25]. Notwithstanding, a central issue surrounding TRPP2 is how much TRPP2 channel activity depends on the presence of PKD1. In a lipid bilayer system, recombinant TRPP2 can reconstitute cation channel activity in the absence of PKD1 [34]. Hanaoka et al. [35] provided a functional substratum for the well-established demonstration

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that PKD1 and TRPP2 interact physically via their C-termini and form heteromeric complexes in vivo [12]. Coexpression of PKD1 and TRPP2 in CHO cells as well as in sympathetic neurons promotes the translocation of TRPP2 to the plasma membrane and generates a nonselective cation channel with perm-selectivity and sensitivity to diand trivalent cations very similar to those of homomeric TRPP2 (Figure 14.1) [35,36]. The channel activity is not observed when C-terminal interaction between PKD1 and TRPP2 is not allowed, implying that coassembly of PKD1 and TRPP2 is required for the targeting/retention of TRPP2 to the plasma membrane. The PKD1/TRPP2 complex has been reconstituted at the cell surface of sympathetic neurons [36]. In this model, TRPP2 is likely to act as the ion-translocating component of the polycystin complex because the pharmacological and permeation properties of the PKD1/TRPP2 channel complex resemble those of recombinant homomeric TRPP2 and because homomeric PKD1 cannot form an ion channel by itself [37]. Within the complex, PKD1 and TRPP2 have reciprocal “stabilizing” effects on each other’s function [34,36]. Indeed, the association of PKD1 appears to repress the constitutive activity of TRPP2 [36], which would be detrimental to the cell if uncontrolled. Conversely, TRPP2 binding to PKD1 represses PKD1’s ability to constitutively activate G proteins possibly by steric/competitive interaction among the different PKD1-binding partners [36,37]. These data favor the view that besides its ion channel function, TRPP2 also regulates the downstream effects of PKD1 on its target effectors and genes. Therefore, the balance between TRPP2 and PKD1 expression, which is manifestly disrupted in ADPKD, may play a critical role in normal PKD1–TRPP2 signaling. TRPP2 repression of G-protein activation by PKD1 has been conﬁrmed in the case of PKD1-mediated NFAT (nuclear factor of activated T-cells) activation [38]; more recently, TRPP2 has been shown to impair the nuclear translocation of the PKD1 C-terminus by binding to, and sequestering of, the latter (Figure 14.1) [39]. TRPP2, in conjunction with PKD1, has been shown to regulate messages to the nucleus by preventing the pro-proliferative helix-loop-helix protein Id2 from entering the nucleus (Figure 14.1) [65]. Id2 is known to associate with E proteins and blocks their ability to turn on growth-suppressive genes. Id2 is normally prevented from translocating to the nucleus through its association with the serine-phosphorylated C-terminal domain of TRPP2, which is promoted by PKD1. These data predict that loss-of-function mutations in either TRPP2 or PKD1, or disruption of their functional interaction, cause Id2 to enter the nucleus and turn off growthsuppressive genes, making a case for the involvement of such a mechanism in the pathogenesis of ADPKD.

ACTIVATION OF THE PKD1–TRPP2 COMPLEX That PKD1 and TRPP2 are interacting partners within a heteromultimeric polycystin complex has been intuited from the observation that mutations in PKD1 or TRPP2 produce virtually identical clinical presentations, irrespective of the causative gene [15]. Reconstituted PKD1–TRPP2 complexes can be activated by applying antibodies directed against the extracellular REJ domain of PKD1 [36]. This activates bidirectional signaling events, concordantly enhancing TRPP2 activity and

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stimulating heterotrimeric G-protein pathways. Thus, PKD1 and TRPP2 form functionally associated subunits of a receptor-ion channel signaling complex in which PKD1 acts as a “receptor/regulator” that controls TRPP2 activity and G proteins (Figure 14.1) [3,34,36]. The activation of TRPP2 and G proteins appears to proceed through a structural rearrangement of the polycystin complex that requires both proteins to have intact C-termini. This mechanism may mimic a yet-to-be-determined (mechanical? ligand?) extracellular signal that activates the polycystin complex. This proposed mechanism may be paradigmatic for the function of other polycystin orthologues in a variety of tissues [3]. For example, PKD1 signaling has fascinating functional parallels with the acrosome reaction (AR) in sea urchin spermatozoa, a prerequisite for sperm-egg fusion [40]. AR requires the activation of suREJ1 or suREJ3, two PKD1 orthologues harboring REJ modules; each binds components of the egg jelly [41]. Importantly, antibodies directed against the REJ domain of suREJs induce the AR by opening Ca2+-permeant channels [42]. Recently, suREJ3 has been shown to physically bind to the sea urchin sperm orthologue of TRPP2 in the acrosome plasma membrane [43], raising the possibility that suTRPP2 may be involved in the Ca2+-regulated AR. Collectively, these ﬁndings add further weight to the primary importance of the REJ domain in activating the polycystin complex. Other early evidence in support of the idea that PKD1 and TRPP2 form heteromultimeric complexes came from studies of C. elegans orthologues of ADPKD genes, lov-1 (location of vulva) and pkd-2 [44]. Lov-1, however, has an unrelated extracellular domain to PKD1 in that it lacks the REJ domain and harbors instead several mucin domains [3]. Mutation analysis shows identical male sensory behavioral defects in single or double lov-1 and pkd-2 mutants, indicating that both proteins act together in a single sensory pathway necessary for normal mating behavior [45]. Lov-1 and pkd-2 concentrate in cilia and cell bodies of male-speciﬁc sensory neurons, consistent with their functions as associated subunits of a receptor-ion channel complex involved in mechanosensitive signaling.

TRPP2: A BONA FIDE MECHANOSENSITIVE CHANNEL? Recent studies have provided evidence that PKD1 and TRPP2 colocalize in primary cilia of renal epithelial cells, where they may function in transducing sensory information, such as shear ﬂuid stress [23]. The primary cilium of renal epithelial cells is a solitary nonmotile structure of a few micrometers that arises from the basal body or centriole and projects into the lumen of the tubule. Its central role in cyst formation has been suggested from the primary observation that defects in proteins necessary for the assembly or function of primary cilia such as cystin, polaris, inversin, and kinesin-II cause polycystic kidney diseases. The cilium is proposed to serve as a ﬂow sensor because it can reversibly bend in response to ﬂuid ﬂow rates comparable to those observed in renal tubules and because it was shown to be essential for Madin-Darby canine kidney (MDCK) cells’ ability to sense ﬂow, because deciliated cells are irresponsive to changes in ﬂow rate [46,47]. Fluid shear-force bending of the cilium causes Ca2+ inﬂux through mechanically

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sensitive channels. Although it is not known whether these mechanosensitive channels reside at the base of cilia or throughout the cilium membrane, one could predict that they localize in the intervening bilayer regions that increase in tension during cilium bending. A calcium signal through mechanically activated channels is then ampliﬁed by Ca2+ release from IP3R stores in MDCK cells and spreads to neighboring cells through gap junctions [46]. These data conﬂict, however, with a study in murine embryonic kidney epithelial (MEK) cells, where ryanodine receptors instead of IP3Rs have been implicated in Ca2+ ampliﬁcation [23]. Thus, the cilium acts as the mechanosensor of changes in laminar ﬂuid ﬂow and transduces stimulus energy into change in membrane permeability. In this model, the PKD1–TRPP2 polycystin complex may be envisioned as a mechanotransducer, which is used to signal relevant intratubular information such as ﬂow rates, directing attention to the regulation of Ca2+ inﬂux as a crucial misstep that initiates cystogenesis. However, whether the PKD1–TRPP2 complex is mechanosensitive still awaits direct experimental evidence. Recently, TRPP2 has been shown to play a central role in establishing the leftright (LR) asymmetry of visceral organs [48], which occurs during early embryonic development when the nodal gene, initially expressed throughout the node, becomes limited to the left margin of the node. The node is a triangular-shaped structure at the distal tip of ∼E7 embryos, consisting of endodermally derived cells, each carrying a single cilium on their apical surface. The monocilia located at the center of the node express dynein, a microtubular motor protein, and are motile, producing rotational movement that creates a leftward ﬂuid ﬂow across the node. This leftward nodal ﬂow is critical for the sideness of asymmetric gene expression because mice with immotile cilia develop laterality defects. In contrast, monocilia located in the periphery of the node are immobile (lacking dynein) and act as sensors of directional nodal ﬂow by generating an asymmetric Ca2+ signal [48]. TRPP2 is expressed in both motile and immotile monocilia, yet a perinodal Ca2+ signal is absent in TRPP2−/− mice embryos, suggesting that TRPP2 functions as a mechanotransducer in immotile monocilia and transduces leftward nodal ﬂow into an increase in Ca2+ at the left border of the node. This function would be key for the establishment of a morphogenic gradient at the embryonic node and consistent with the observation that targeted disruption in TRPP2 causes situs inversus in addition to the hallmark cardiac and kidney defects [49]. The lack of laterality defects in PKD1 knockout embryos correlates with the absence of PKD1 in cilia [50], favoring the idea that TRPP2 may be involved in mechanosensation in the absence of PKD1. In the same vein, the orthologue of TRPP2 encoded by the amo gene (almost there) in Drosophila melanogaster is localized to the distal tip of the sperm ﬂagella and, apparently in the absence of the PKD1 orthologue, plays a critical role for directional movement inside the female reproductive tract [51,52]. This suggests that TRPP2 in Drosophila sperm is part of a signaling pathway involved in detecting directional cues that are necessary for entry into the female storage organs, perhaps supporting a common role for Ca2+-dependent TRPP2 signals in both motile and immotile axonemal-based structures. Despite the widespread utilization of mechanosensitive channels in a variety of physiological processes including the detection of touch, hearing function, blood

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pressure control, and osmotic pressure, little is known about the molecular structure and organization of vertebrate mechanotransducers. In this respect, the recent identiﬁcation of TRP channels as core components of mechanoreceptors in C. elegans, Drosophila melanogaster, and vertebrates may offer clues to the conservative mechanoreceptive structural elements of mechanotransducers [53,54]. In the fruit ﬂy, mechanoelectrical responses in bristle sensory neurons occur rapidly upon deﬂection of the bristle hair shaft and result from the opening—among others—of the NOMPC channel, a member of the TRPN subfamily. NOMPC has a particularly long intracellular amino-terminal tail harboring 29 ankyrin repeats, which are considered to anchor the channel to the cytoskeleton and may mediate the protein–protein interaction of a tethered mechanism that might be required for mechanical gating. The nematode C. elegans senses nose touch by stimulating ciliated nociceptive sensory neurons, which detect, among others, mechanical and osmotic stimuli. The OSM-9 channel is thought to be part of the mechanosensitive channel because OSM9 mutants are defective in osmotic avoidance and in sensitivity to nose touch. OSM-9 is a homologue to members of the TRPV channels, with three ankyrin-repeat domains at its amino-terminal intracellular domain. TRPV2 and TRPV4, its invertebrate TRP counterparts, have multiple ankyrin-repeat domains and are implicated in vertebrate mechanosensation in that they can sense membrane stretch [55] and hypo-osmotic stress [56], respectively. With regard to mechanical gating, it is noteworthy that TRPV4 requires the amino-terminal domain with the three ankyrinrepeats to sense physical challenges [57]. More recently, Corey et al. [58] have shown that TRPA1 (also called ANKTM1), which harbors 17 ankyrin domains, constitutes or is a component of the mechanosensitive transduction channel of vertebrate hair cells. Although the mechanism of activation by mechanical force is not yet established, an ankyrin repeat has been hypothesized to form a springlike gating structure, consistent with a “tethered channel” model [4]. Neither PKD1 nor TRPP2 display ankyrin-repeats that would allow tight interactions between the channel complex and the cytoskeleton. TRPP2, however, has been shown to connect indirectly with the actin cytoskeletal network, though it remains to be shown whether these actin-based elements play a role in cilium mechanotransduction, given that the cilium is primarily the domain of microtubules rather than actin ﬁlaments [3]. On this last issue, the extracellular domain of PKD1 has been shown to display a dynamic extensibility whereby its length might be regulated through unfolding/refolding its Ig-like domains [59]. Although these mechanical properties of PKD1 are important in the context of mechanosensation, they seem more appropriate to provide structural support in cell–cell or cell–matrix interactions at basolateral membranes than mechanosensation in the solitary cilium. At this point, the available information does not entirely support the candidacy of a PKD1–TRPP2 complex as the core component of the primary cilium’s mechanosensitive apparatus. This would seem ample reason to consider alternative models in which mechanical force is transmitted indirectly to the protein complex or via an auxiliary subunit. On the one hand, this may imply that the TRPP2 channel acts as a nonmechanosensitive ampliﬁer of a true mechanically gated channel, with cytosolic Ca2+ acting as a suitable activator of TRPP2. A critical issue to be

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established is the polymodal nature of TRPP2 regulation (i.e., whereby channel opening is not only dependent on mechanical stimuli but also modulated by PKD1, Ca2+, H+, and phosphorylation). Thus, the basic distinction between physical and chemical mechanisms of PKD1–TRPP2 mechanotransduction is not yet made. On the other hand, it is also conceivable that TRPP2 coassembles with other TRP channels to form a mechanosensitive channel as many invertebrate TRP-related channels do. A tantalizing link points to TRPV4, because it is expressed in renal epithelial cells, particularly in the distal nephron and collecting ducts, which are ﬂow-sensitive segments [60]. In this regard, preliminary evidence by Walz’s group indicate that TRPV4 and TRPP2 functionally interact and colocalize in the primary cilium [61]. TRPC1 has also been recently found to be a component of the vertebrate mechanosensitive cation channel [62]. In contrast to TRPA1, TRPC1 is gated by tension developed in the lipid bilayer. Interestingly, TRPC1 is known to interact with TRPP2 in expression systems [63] and to form functional heterotetramers with TRPP2 [64], suggesting that TRPC1 as well may contribute to the mechanosensory TRPP2 apparatus (Figure 14.1).

CONCLUDING REMARKS In the past ﬁve years, considerable progress has been achieved in evaluating the distribution, pathophysiology, and functional characteristics of TRPP proteins. Most notably, TRPP2 channels have been shown to trafﬁc to different subcellular compartments and to display speciﬁc subcellular functions. In renal primary cilia, TRPP2 interacts with PKD1, a process that may be essential functionally for the regulation of mechanosensation and the cell cycle. In the ER, TRPP2 serves as an intracellular Ca2+ release channel. It would be important to identify ligands for PKD1 that affect the function of the PKD1–TRPP2 complex and to identify the factors interacting with TRPP2 that determine and regulate its compartment-speciﬁc functions. These studies will contribute not only to a better understanding of TRPP physiological functions but also to the development of new strategies for targeted therapeutic intervention.

ACKNOWLEDGMENTS This work was supported by the Centre National de la Recherche Scientiﬁque (CNRS), and by grants from the Agence Nationale de la Recherche (ANR Cardiovasculaire, obésité et diabète, N° ANR-05-PCOD-029-02), la Fondation Schlumberger pour l’Education et la Recherche, the French Ministère délégué à la Recherche (ACI Jeune Chercheurs, N° 5294), and a fellowship from the French Research Ministry and the University of Méditerranée Marseille.

REFERENCES 1. Nilius, B. and Voets, T. TRP channels: a TR(I)P through a world of multifunctional cation channels. Pﬂügers Arch. 451, 1, 2005.

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2. Montell, C. et al. A uniﬁed nomenclature for the superfamily of TRP cation channels. Mol. Cell 9, 229, 2002. 3. Delmas, P. Polycystins: from mechanosensation to gene regulation. Cell 118, 145, 2004. 4. Delmas, P. Polycystins: polymodal receptor/ion-channel cellular sensors. Pﬂügers Arch. 451, 264, 2005. 5. Igarashi, P. and Somlo, S. Genetics and pathogenesis of polycystic kidney disease. J. Am. Soc. Nephrol. 13, 2384, 2002. 6. Nauli, S.M. and Zhou, J. Polycystins and mechanosensation in renal and nodal cilia. Bioessays 26, 844, 2004. 7. Montell, C. The TRP superfamily of cation channels. Sci. STKE, re3, 2005. 8. Chen, X.Z. et al. Polycystin-L is a calcium-regulated cation channel permeable to calcium ions. Nature 401, 383, 1999. 9. Köttgen, M. et al. Trafﬁcking of TRPP2 by PACS proteins represents a novel mechanism of ion channel regulation. EMBO J. 24, 705, 2005. 10. Delmas, P. et al. Polycystins, calcium signaling, and human diseases. Biochem. Biophys. Res. Commun. 322, 1374, 2004. 11. Qian, F. et al. Cleavage of polycystin-1 requires the receptor for egg jelly domain and is disrupted by human autosomal-dominant polycystic kidney disease 1-associated mutations, Proc. Natl. Acad. Sci. USA 99, 16981, 2002. 12. Newby, L.J. et al. Identiﬁcation, characterization, and localization of a novel kidney polycystin-1–polycystin-2 complex. J. Biol. Chem. 277, 20763, 2002. 13. Murakami, M. et al. Genomic organization and functional analysis of murine PKD2L1. J. Biol. Chem. 280, 5626, 2005. 14. Li, A., Tian, X., Sung, S.W., and Somlo, S. Identiﬁcation of two novel polycystic kidney disease-1-like genes in human and mouse genomes. Genomics 81, 596, 2003. 15. Sutters, M. and Germino, G.G. Autosomal dominant polycystic kidney disease: molecular genetics and pathophysiology. J. Lab. Clin. Med. 141, 91, 2003. 16. Gogusev, J. et al. Molecular cytogenetic aberrations in autonomal dominant polycystic kidney disease tissue. J. Am. Soc. Nephrol. 14, 359, 2003. 17. Ong, A.C. Polycystin expression in the kidney and other tissues: complexity, consensus and controversy. Exp. Nephrol. 8, 208, 2000. 18. Cai, Y. et al. Identiﬁcation and characterization of polycystin-2, the PKD2 gene product. J. Biol. Chem. 274, 28557, 1999. 19. Chen, X.Z. et al. Transport function of the naturally occurring pathogenic polycystin2 mutant, R742X. Biochem. Biophys. Res. Commun. 282, 1251, 2001. 20. Luo, Y. et al. Native polycystin 2 functions as a plasma membrane Ca2+-permeable cation channel in renal epithelia. Mol. Cell Biol. 23, 2600, 2003. 21. Vassilev, P.M. Polycystin-2 is a novel cation channel implicated in defective intracellular Ca2+ homeostasis in polycystic kidney disease. Biochem. Biophys. Res. Commun. 282, 341, 2001. 22. Gonzalez-Perrett, S. Polycystin-2, the protein mutated in autosomal dominant polycystic kidney disease (ADPKD), is a Ca2+-permeable nonselective cation channel, Proc. Natl. Acad. Sci. USA 98, 1182, 2001. 23. Nauli, S.M. Polycystins 1 and 2 mediate mechanosensation in the primary cilium of kidney cells. Nat. Genet. 33, 129, 2003. 24. Yoder, B.K., Hou, X., and Guay-Woodford, L.M. The polycystic kidney disease proteins, polycystin-1, polycystin-2, polaris and cystin, are co-localized in renal cilia. J. Am. Soc. Nephrol. 13, 2508, 2002.

Activation Mechanisms and Functional Roles of TRPP2 Cation Channels

201

25. Foggensteiner, L. et al., Cellular and subcellular distribution of polycystin-2, the protein product of the PKD2 gene. J. Am Soc. Nephrol. 11, 814, 2000. 26. Rundle, D.R., Gorbsky, G.J., and Tsiokas, L. PKD2 interacts and co-localizes with mDia1 to mitotic spindles of dividing cells: role of mDia1 in PKD2 localization to mitotic spindles. J. Biol. Chem. 279, 29728, 2004. 27. Hidaka, S., Könecke, V., Osten, L., and Witzgall, R. PIGEA-14, a novel coiled-coil protein affecting the intracellular distribution of polycystin-2. J. Biol. Chem. 279, 35009, 2004. 28. Köttgen, M. and Walz, G. Subcellular localization and trafﬁcking of polycystins. Pﬂügers Arch. 451, 286, 2005. 29. Koulen, P. Polycystin-2 is an intracellular calcium release channel. Nat. Cell Biol. 4, 191, 2002. 30. Li, Q. Liu, Y., Zhao, W. and Chen, X.Z. The calcium-binding EF-hand in polycystinL is not a domain for channel activation and ensuing inactivation. FEBS Lett. 516, 270, 2002. 31. Cai, Y. et al. Calcium dependence of polycystin-2 channel activity is modulated by phosphorylation at Ser812. J. Biol. Chem. 279, 19987, 2004. 32. Qian, Q. et al. Pkd2 haploinsufﬁciency alters intracellular calcium regulation in vascular smooth muscle cells. Hum. Mol. Genet. 12, 1875, 2003. 33. Li, Y. et al. Polycystin 2 interacts with type I inositol 1,4,5-triphosphate receptor to modulate intracellular Ca2+ signaling. J. Biol. Chem. 280, 41298, 2005. 34. Xu, G.M. et al., Polycystin-1 activates and stabilizes the polycystin-2 channel. J. Biol. Chem. 278, 1457, 2003. 35. Hanaoka, K. et al. Co-assembly of polycystin-1 and -2 produces unique cationpermeable currents. Nature 408, 990, 2000. 36. Delmas, P. et al. Gating of the polycystin ion channel signaling complex in neurons and kidney cells. FASEB J. 18, 740, 2004. 37. Delmas, P. et al. Constitutive activation of G-proteins by polycystin-1 is antagonized by polycystin-2. J. Biol. Chem. 277, 11276, 2002. 38. Puri, S. et al. Polycystin-1 activates the calcineurin/NFAT (nuclear factor of activated T-cells) signaling pathway. J. Biol. Chem. 279, 55455, 2004. 39. Chauvet, V. et al. Mechanical stimuli induce cleavage and nuclear translocation of the polycystin-1 C-terminus. J. Clin. Invest. 114, 1433, 2004. 40. Mengerink, K.J., Moy, G.W., and Vacquier, V.D. suREJ proteins: new signalling molecules in sea urchin spermatozoa. Zygote 8, S28, 2000. 41. Hirohashi, N. and Vacquier, V.D. High molecular mass egg fucose sulfate polymer is required for opening both Ca2+ channels involved in triggering the sea urchin sperm acrosome reaction. J. Biol. Chem. 277, 1182, 2002. 42. Moy, G.W. et al. The sea urchin sperm receptor for egg jelly is a modular protein with extensive homology to the human polycystic kidney disease protein, PKD1. J. Cell Biol. 133, 809, 1996. 43. Neill, A.T., Moy, G.W., and Vacquier, V.D. Polycystin-2 associates with the polycystin-1 homolog, suREJ3, and localizes to the acrosomal region of sea urchin spermatozoa. Mol. Reprod. Dev. 67, 472, 2004. 44. Barr, M.M. and Sternberg, P.W. A polycystic kidney-disease gene homologue required for male mating behaviour in C. elegans. Nature 401, 386, 1999. 45. Barr, M.M. et al. The Caenorhabditis elegans autosomal dominant polycystic kidney disease gene homologs lov-1 and pkd-2 act in the same pathway, Curr. Biol. 11, 1341, 2001.

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46. Praetorius, H.A. and Spring, K.R. Bending the MDCK cell primary cilium increases intracellular calcium. J. Membr. Biol. 184, 71, 2001. 47. Praetorius, H.A. and Spring, K.R. Removal of the MDCK cell primary cilium abolishes ﬂow sensing. J. Membr. Biol. 191, 69, 2003. 48. McGrath, J. et al. Two populations of node monocilia initiate left-right asymmetry in the mouse. Cell, 114, 61, 2003. 49. Pennekamp, P. et al. The ion channel polycystin-2 is required for left-right axis determination in mice. Curr. Biol. 12, 938, 2002. 50. Karcher, C. et al. Lack of laterality phenotype in Pkd1 knock-out embryos correlates with the absence of polycystin-1 in nodal cilia. Differentiation 73, 425, 2005. 51. Gao, Z., Ruden, D.M., and Lu, X. PKD2 cation channel is required for directional sperm movement and male fertility. Curr. Biol. 13, 2175, 2003. 52. Watnick, T.J. et al. A ﬂagellar polycystin-2 homolog required for male fertility in Drosophila. Curr. Biol. 13, 2179, 2003. 53. Pedersen, S.F., Owsianik, G., and Nilius, B. TRP channels: an overview. Cell Calcium 38, 233, 2005. 54. O’Neil, R.G. and Heller, S. The mechanosensitive nature of TRPV channels. Pﬂügers Arch. 451, 193, 2005. 55. Muraki, K. et al. TRPV2 is a component of osmotically sensitive cation channels in murine aortic myocytes. Circ. Res. 93, 829, 2003. 56. Alessandri-Haber, N. et al. Hypotonicity induces TRPV4-mediated nociception in rat. Neuron 39, 497, 2003. 57. Liedtke, W. et al. Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor. Cell 103, 525, 2000. 58. Corey, D.P. et al. TRPA1 is a candidate for the mechanosensitive transduction channel of vertebrate hair cells. Nature 432, 723, 2004. 59. Qian, F. et al. The nanomechanics of polycystin-1 extracellular region. J. Biol. Chem. 280, 40723, 2005. 60. Tian, W. et al. Renal expression of osmotically responsive cation channel TRPV4 is restricted to water-impermeant nephron segments. Am. J. Physiol. 287, F17, 2004. 61. Köttgen, M. et al. Polycystin-2 and TRPV4 form a functional heteromultimeric complex that might act as a cilial mechanosensor. J. Am. Soc. Nephrol. 16, TH-FC116, 2005. 62. Maroto, R. et al. TRPC1 forms the stretch-activated cation channel in vertebrate cells. Nature Cell Biol. 7, 179, 2005. 63. Tsiokas, L. et al. Speciﬁc association of the gene product of PKD2 with the TRPC1 channel. Proc. Natl. Acad. Sci. USA 9, 3934, 1999. 64. Delmas, P. Assembly and gating of TRPC channels in signalling microdomains. Novartis Found Symp. 258, 75, 2004. 65. Li, X. et al. Polycystin-1 and polycystin-2 regulate the cell cycle through the helix-loophelix inhibitor Id2. Nature Cell Biol. 7, 1202, 2005.

15

The Ca2+-Activated TRP Channels: TRPM4 and TRPM5 Emily R. Liman University of Southern California

CONTENTS Cloning, Distribution, and Structure of TRPM4 and TRPM5 .............................204 Ion Permeability, Unitary Properties, and Speciﬁc Blockers of TRPM4 and TRPM5 .........................................................................................204 Sensitivity of TRPM4 and TRPM5 to Intracellular Ca2+ .....................................205 Activation of TRPM4 and TRPM5 by Voltage.....................................................206 Regulation of TRPM4 and TRPM5 by PIP2 .........................................................207 Function of TRPM4 and TRPM5..........................................................................208 Note........................................................................................................................210 References..............................................................................................................210

Ca2+-activated nonselective cation channels were ﬁrst described in the early 1980s [1,15,31], shortly after the patch-clamp recording technique was developed [5]. Patch-clamp recording, and in particular the excised-patch recording conﬁguration, made it possible to rapidly and reproducibly change the intracellular milieu of cells and therefore to discover intracellular activators and modulators of ion channels. In this context, intracellular Ca2+ at micromolar concentrations was found to activate ion channels that were equally permeable to Na+ and K+ ions, and these ion channels were detected in a variety of cell types. The Ca2+-activated nonselective channels found in cultured heart cells were proposed to play a role in modulating the duration of the action potential, whereas those in pancreatic acinar cells were proposed to play a role in exocrine secretion [1,15]. However, without speciﬁc pharmacological or genetic tools, a clear attribution of function was difﬁcult. In 2002 this situation changed with the cloning and functional expression of the ﬁrst Ca2+-activated nonselective cation channel, TRPM4 [9]. Following this discovery, a second Ca2+activated cation channel, TRPM5, was characterized [6,12,22,24,32]. Our understanding of the biological signiﬁcance of these channels, their functional regulation, and their structural properties has since proceeded at a rapid pace.

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CLONING, DISTRIBUTION, AND STRUCTURE OF TRPM4 AND TRPM5 TRPM4 and TRPM5 are members of the TRPM subfamily of transient receptor potential ion channels. The ﬁrst member of the TRPM subfamily of ion channels, melastatin (TRPM1), was identiﬁed as a gene that was downregulated in mouse melanoma cell lines [2]. Based on similarity to TRPM1, a partial sequence of TRPM4 was detected in the database of human-expressed sequence tags and used to identify full-length cDNAs with two possible splice variants encoding proteins of 1,040 (TRPM4a) [30] and 1,214 (TRPM4b) [9] amino acids. Subsequently, only TRPM4b has been shown to be functional and therefore the term TRPM4 will be used to refer to this splice variant. TRPM4’s message is expressed widely and is at its highest levels in the heart, prostate, colon [30], placenta, and pancreas [9]. No localization of the mRNA or protein has yet been reported within these tissues. TRPM5 was initially identiﬁed by homology to TRPM1 [4,23] and was later found by two independent groups to be a key component of mammalian taste cells [22,32]. Presently only a single splice variant has been reported. In contrast to TRPM4, TRPM5 mRNA is expressed in a highly restricted manner, and among major tissues of the body it is found only at high levels in the tongue, small intestine, and stomach [22]. Lower levels of TRPM5 may be expressed in other tissues [23]. TRPM4 and TRPM5, which are structurally more related to each other than to other TRP channels, show ~40 percent amino acid identity. Based on hydropathy analysis and homology to voltage-activated ion channels, both have been proposed to contain six transmembrane-spanning regions and to assemble as tetramers. The cytoplasmic N- and C-termini of TRPM4 and TRPM5 contain a number of potential protein–protein binding sites: the C-termini of both channels contain predicted coiled-coil domains and N- and C-termini of TRPM4 contain calmodulin-binding sites [19]. The pore of TRPM4 appears to be located between the ﬁfth and sixth transmembrane domains, an assignment that is supported by the observation that mutations within this region alter ion selectivity [17]. Moreover, a histidine residue in the C-terminal portion of this region mediates an external proton block of TRPM5, supporting the contention that this region forms the outer pore vestibule [13].

ION PERMEABILITY, UNITARY PROPERTIES, AND SPECIFIC BLOCKERS OF TRPM4 AND TRPM5 Both TRPM4 and TRPM5 are robustly activated by intracellular Ca2+, making it possible to study their function properties in detail with patch-clamp recording [6,9,12,16,24]. These studies have revealed a number of similarities and differences between the two channels that will be helpful in classifying native channels and in understanding structure-function relationships. Both channels have a unitary conductance of ~25 pS [6,9,12,24], but whereas the openings of TRPM4 channels are long-lived (several hundred milliseconds; e.g., see reference 33), openings of TRPM5 channels are transient (e.g., see reference 12). Both channels are permeable to monovalent cations and impermeable to divalent cations [6,9,12,24].

The Ca2+-Activated TRP Channels: TRPM4 and TRPM5

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At present only a few “speciﬁc” blockers of TRPM4 and TRPM5 channels have been identiﬁed. TRPM4 channels are blocked by micromolar concentrations of intracellular ATP and other adenine nucleotides [20], whereas TRPM5 channels are insensitive to these small molecules. TRPM5 channels are blocked by external pH of 6.0 or lower [13], whereas TRPM4 channels are insensitive to acid pH. Based on these characteristics, one can assign the likely genetic basis for native channel activity; for example, based on the above criteria, the Ca2+-activated channels expressed in pheromone-sensing cells of the vomeronasal organs, which show long openings and are blocked by adenine nucleotides, are likely encoded by TRPM4 [10].

SENSITIVITY OF TRPM4 AND TRPM5 TO INTRACELLULAR CA2+ One of the most important ways in which TRPM4 and TRPM5 channels differ is in sensitivity to intracellular Ca2+. In excised inside-out patches immediately upon patch excision, the EC50 for activation by Ca2+ of mTRPM4 is 100–200 μM [27,33], and the EC50 for activation by Ca2+ of mTRPM5 is 20–30 μM [12,27] (Figure 15.1). Sensitivity to Ca2+ decreases for both channels by ~four-fold following patch excision and exposure to Ca2+; the EC50 for activation by Ca2+ after desensitization is ~500 μM and ~80 μM for TRPM4 and TRPM5, respectively [12,33]. Somewhat mysteriously, TRPM4 and TRPM5 currents in whole-cell recording mode are ~10–50 times more sensitive to Ca2+, and both can be maximally activated by concentrations of Ca2+ as low as 1 μM (e.g., see reference 27). It is not known why

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FIGURE 15.1 Ca2+-dependent activation of TRPM4 and TRPM5 currents. (A,B) Responses of TRPM4 and TRPM5 currents to increasing concentrations of Ca2+. The lower panels show currents recorded from the same patches after desensitization. (C) Dose-response relations of TRPM4 and TRPM5 currents to activation by Ca2+. Modiﬁed from references 12 and 33 with permission.

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there is such a large discrepancy in Ca2+ sensitivity between excised-patch and wholecell recording modes. One possibility is that a factor is lost upon patch excision that controls sensitivity (see below).

ACTIVATION OF TRPM4 AND TRPM5 BY VOLTAGE In the presence of an activating concentration of intracellular Ca2+, depolarization strongly increases the opening of TRPM4 and TRPM5 channels [6,12,16,26]. This is apparent in the strong outward rectiﬁcation of TRPM4 and TRPM5 currents in response to voltage ramps, despite the fact that the underlying channels display a linear I–V relation [6,12,16,24,26]. In response to depolarizing voltage steps, both TRPM4 and TRPM5 currents show time-dependent relaxation, reﬂecting the opening of channels (Figure 15.2B). Voltage-dependent activation has also been reported for TRPM8 and TRPM4 and thus may be a common feature of TRPM channels [6,16,21,25]. It has been hypothesized that the weak voltage dependence of these channels allows their gating to be easily modulated [21], a hypothesis that is supported by work on cold regulation of TRPM8, TRPV1, and TRPM5 [26,29], decavanadate modulation of TRPM4 [18], and PI(4,5)P2 regulation of TRPM4 and TRPM8 [25,33] (see below). The structural mechanism of voltage sensing of these TRP channels is not presently understood, but the presence of several positively charged residues in the fourth transmembrane of these channels suggests that this may be the voltage sensor [7,21].

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FIGURE 15.2 TRPM5 currents are Ca2+ and voltage dependent. (A) In whole-cell recording mode, 40 μM Ca2+ in the pipette elicited a large rectifying current in a HEK293 cell expressing TRPM5. Recording began shortly after break in to the whole-cell mode. Inset shows the current in response to a ramp depolarization (1 V/s). (B) TRPM5 currents in response to a family of step depolarizations and the resulting I–V relationship for the peak current at each voltage. Steps are to 0–100 mV from a holding potential of –80 mV with repolarization to –50 mV. Modiﬁed from reference 12 with permission.

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REGULATION OF TRPM4 AND TRPM5 BY PIP2 A consistent observation is that TRPM4 and TRPM5 currents run down following activation, regardless of whether the currents are measured in whole-cell, perforated-patch, or excised-patch recording modes (Figure 15.2A and Figure 15.3). In excised patches, this rundown is associated with a decrease in the Ca2+ sensitivity of the channels, and thus can be formally considered desensitization [12,33]. What is the basis for desensitization? Recent evidence indicates that it is at least in part due to hydrolysis of membrane phophoinosides, most likely by Ca2+-activated membraneassociated PLCs [28]. The evidence for this conclusion includes the demonstration that (1) both TRPM5 and TRPM4 currents can be rescued from desensitization by PI(4,5)P2 [12,33] (Figure 15.3C); (2) TRPM4 currents can be rescued from desensitization by intracellular MgATP at concentrations that activate lipid kinases and are expected to restore PI(4,5)P2 levels, and this effect can be blocked by lipid kinase

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FIGURE 15.3 ATP and PI(4,5)P2 restore TRPM4 currents from desensitization. Inward currents evoked in response to 100 μM cytoplasmic Ca2+ were recorded from inside-out patches from TRPM4-expressing ChoK1 cells (Vm = –80 mV). (A) Following desensitization, TRPM4 currents can be recovered by exposure to MgATP (2 mM) but not by free ATP (4 mM). (B) Rundown of the TRPM4 current is accelerated by exposure to the PI(4,5)P2 scavenger, poly-L-lysine. (C) In a patch with fast rundown, DiC8 PI(4,5)P2 (10 μM) restores the mTRPM4 current to its initial magnitude. Modiﬁed from reference 33 with permission.

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inhibitors [19,33] (Figure 15.3A); (3) desensitization of TRPM4 is promoted by depletion of PI(4,5)P2 with polyamines (Figure 15.3B). At present the mechanism and structural determinants for PI(4,5)P2 regulation of TRPM4 and TRPM5 is not known. Detailed study shows that PI(4,5)P2 changes the voltage-dependent gating of TRPM4 channels, promoting channel activation at negative membrane potentials by stabilizing the channels in the “voltage-activated” state [33]. This mechanism is similar to one proposed to explain PI(4,5)P2 regulation of the structurally related, cold-activated channel TRPM8 [25]. Sensitivity to PI(4,5)P2 is conferred by positively charged residues in the TRP domain of TRPM8, a loosely conserved sequence of 25 amino acids located adjacent to the sixth transmembrane domain [25]. It is not known whether this region also mediates interaction of TRPM4 with PI(4,5)P2.

FUNCTION OF TRPM4 AND TRPM5 Insight into the function of TRPM5 came with the discovery that its expression is largely restricted to taste cells and the GI tract [22,32]. Taste consists of ﬁve distinct modalities of which three—bitter, sweet, and umami—are mediated by G-proteincoupled receptors. These receptors activate a signaling cascade of which several downstream targets have been identiﬁed, including Gα13, gustducin, and PLCβ2 [11,14]. TRPM5 is coexpressed with all three types of G-protein-coupled receptors as well as with their downstream signaling components [22,32], suggesting that it acts in the pathway for bitter, sweet, and umami taste transduction. In support of this idea, knockout of TRPM5 or PLCβ2 severely impairs the ability of mice to detect bitter, sweet, and umami tastes [32]. Based on the present genetic and physiological data, a preliminary model for transduction of bitter, sweet, and amino acid tastes can be proposed. In this model, receptor activation of PLCβ2 generates IP3, which releases Ca2+ from intracellular stores, and store-released Ca2+ opens TRPM5 channels. Opening of TRPM5 channels leads to a depolarization of the taste cell and transmitter release. Adaptation of taste responses may be partly mediated by the hydrolysis of PI(4,5)P2, which is a cofactor for activation of TRPM5 [12] (Figure 15.4, left). TRPM4’s function has been more difﬁcult to discern and a knockout of TRPM4, which would expedite this task, has not yet been reported. Moreover, TRPM4 mRNA is widely distributed, suggesting that it may play a less speciﬁc functional role than TRPM5. Despite these difﬁculties, there is strong evidence that TRPM4 plays a role in cytokine secretion by T lymphocytes and myogenic constriction of cerebral arteries [3,8]. T lymphocytes respond to stimulation with phyohemagglutinin (PHA) with Ca2+ oscillations that release the cytokine IL-2. Dominant negative suppression of TRPM4 and RNA knockdown of TRPM4 prolongs the Ca2+ response, leading to an increase in IL-2 production [8]. Thus TRPM4 appears to play a role in dampening the response of T cells to PHA. Based on these results, a model has been proposed in which activation of T-cell receptors induces a PLC-signing pathway leading to the depletion of Ca2+ from intracellular stores and the consequent entry of Ca2+ through CRAC channels. Ca2+ entry triggers activation of Ca2+-activated K+ channels,

The Ca2+-Activated TRP Channels: TRPM4 and TRPM5

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T cell signaling

FIGURE 15.4 Models for physiological activation of TRPM5 and TRPM4. (Left) A model for taste transduction. Binding of taste stimuli to G-protein-coupled taste receptors (T1R and T2R) leads to dissociation of the heterotrimeric G protein. βγ subunits of the G protein activate PLC2, which in turn hydrolyzes PIP2 into DAG and IP3. IP3 activates IP3 receptors, which release Ca2+ from intracellular stores. Intracellular Ca2+ opens TRPM5 channels, leading to an inﬂux of Na+ and depolarization of the cell. Modiﬁed from reference 12 with permission. (Right) A model for the generation of Ca2+ oscillations in T cells. The T-cell receptor (TCR) activates a PLC-signaling cascade similar to that in taste cells. In response to depletion of Ca2+ stores, CRAC channels are activated, which conduct Ca2+ into the cell. Ca2+ entry activates Ca2+-activated K+ channels and TRPM4 channels. Differences in the temporal pattern of activation of these two channels leads to oscillations in the membrane potential [8].

the opening of which promotes entry of Ca2+ by increasing the driving force across the plasma membrane. Activation of TRPM4 by Ca2+ entering through CRAC channels causes a depolarization of the membrane potential, which opposes further Ca2+ entry and promotes the opening of V-gated K+ channels. The interplay of these four conductances generates the Ca2+ oscillations observed in response to stimulation of T-cell receptors [8] (Figure 15.4, right). Two models for the functional roles of Ca2+-activated TRPs have been proposed [8,12]. One posits that TRPM5 is activated by Ca2+ released from intracellular stores and that TRPM5 generates the primary electrical response to tastants. The second posits that TRPM4 is activated by Ca2+ entry through CRAC channels and that the primary function of TRPM4 is to dampen this signaling pathway. In this context, it may be signiﬁcant that TRPM4 and TRPM5 differ widely in Ca2+ sensitivity and thus may respond to different Ca2+ signals in cells. For example, the higher sensitivity of TRPM5 to Ca2+ may allow it to respond to global changes in Ca2+ concentration following release of Ca2+ from intracellular stores, whereas TRPM4 might be insensitive to such signals and respond only to entry of Ca2+ through closely opposed membrane Ca2+ channels. A challenge will be to determine the sources of Ca2+ that activate these two channels under physiological conditions and to identify the speciﬁc attributes of each channel that allow it to respond selectively to the appropriate stimulus.

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NOTE After this chapter was submitted, Nilius et al. [34] showed a similar regulation of TRPM4 by PIP2 as we described.

REFERENCES [1] [2]

[3]

[4]

[5]

[6]

[7] [8] [9]

[10] [11] [12] [13]

[14] [15] [16]

[17]

D. Colquhoun, E. Neher, H. Reuter, and C.F. Stevens, Inward current channels activated by intracellular Ca in cultured cardiac cells, Nature 294 (1981) 752–754. L.M. Duncan, J. Deeds, J. Hunter, J. Shao, L.M. Holmgren, E.A. Woolf, R.I. Tepper, and A.W. Shyjan, Down-regulation of the novel gene melastatin correlates with potential for melanoma metastasis, Cancer Res. 58 (1998) 1515–1520. S. Earley, B.J. Waldron, and J.E. Brayden, Critical role for transient receptor potential channel TRPM4 in myogenic constriction of cerebral arteries, Circ. Res. 95 (2004) 922–929. T. Enklaar, M. Esswein, M. Oswald, K. Hilbert, A. Winterpacht, M. Higgins, B. Zabel, and D. Prawitt, Mtr1, a novel biallelically expressed gene in the center of the mouse distal chromosome 7 imprinting cluster, is a member of the Trp gene family, Genomics 67 (2000) 179–187. O.P. Hamill, A. Marty, E. Neher, B. Sakmann, and F.J. Sigworth, Improved patchclamp techniques for high-resolution current recording from cells and cell-free membrane patches, Pﬂügers Arch. 391 (1981) 85–100. T. Hofmann, V. Chubanov, T. Gudermann, and C. Montell, TRPM5 is a voltagemodulated and Ca(2+)-activated monovalent selective cation channel, Curr. Biol. 13 (2003) 1153–1158. Y. Jiang, V. Ruta, J. Chen, A. Lee, and R. MacKinnon, The principle of gating charge movement in a voltage-dependent K+ channel, Nature 423 (2003) 42–48. P. Launay, H. Cheng, S. Srivatsan, R. Penner, A. Fleig, and J.P. Kinet, TRPM4 regulates calcium oscillations after T cell activation, Science 306 (2004) 1374–1377. P. Launay, A. Fleig, A.L. Perraud, A.M. Scharenberg, R. Penner, and J.P. Kinet, TRPM4 is a Ca2+-activated nonselective cation channel mediating cell membrane depolarization, Cell 109 (2002) 397–407. E.R. Liman, Regulation by voltage and adenine nucleotides of a Ca2+-activated cation channel from hamster vomeronasal sensory neurons, J. Physiol. 548 (2003) 777–787. B. Lindemann, Receptors and transduction in taste, Nature 413 (2001) 219-225. D. Liu and E.R. Liman, Intracellular Ca2+ and the phospholipid PIP2 regulate the taste transduction ion channel TRPM5, Proc. Natl. Acad. Sci. USA 100 (2003) 15160–15165. D. Liu, Z. Zhang, and E.R. Liman, Extracellular acid block and acid-enhanced inactivation of the Ca2+-activated cation channel TRPM5 involve residues in the S3–S4 and S5–S6 extracellular domains, J. Biol. Chem. 280 (2005) 20691–20699. R.F. Margolskee, Molecular mechanisms of bitter and sweet taste transduction, J. Biol. Chem. 277 (2002) 1–4. Y. Maruyama and O.H. Petersen, Single-channel currents in isolated patches of plasma membrane from basal surface of pancreatic acini, Nature 299 (1982) 159–161. B. Nilius, J. Prenen, G. Droogmans, T. Voets, R. Vennekens, M. Freichel, U. Wissenbach, and V. Flockerzi, Voltage dependence of the Ca2+-activated cation channel TRPM4, J. Biol. Chem. 278 (2003) 30813–30820. B. Nilius, J. Prenen, A. Janssens, G. Owsianik, C. Wang, M.X. Zhu, and T. Voets, The selectivity ﬁlter of the cation channel TRPM4, J. Biol. Chem. 280 (2005) 22899–22906.

The Ca2+-Activated TRP Channels: TRPM4 and TRPM5 [18] [19]

[20]

[21] [22]

[23]

[24]

[25]

[26]

[27]

[28]

[29]

[30]

[31] [32]

[33]

[34]

211

B. Nilius, J. Prenen, A. Janssens, T. Voets, and G. Droogmans, Decavanadate modulates gating of TRPM4 cation channels, J. Physiol. 560 (2004) 753–765. B. Nilius, J. Prenen, J. Tang, C. Wang, G. Owsianik, A. Janssens, T. Voets, and M.X. Zhu, Regulation of the Ca2+ sensitivity of the nonselective cation channel TRPM4, J. Biol. Chem. 280 (2005) 6423–6433. B. Nilius, J. Prenen, T. Voets, and G. Droogmans, Intracellular nucleotides and polyamines inhibit the Ca2+-activated cation channel TRPM4b, Pﬂügers Arch. 448 (2004) 70–75. B. Nilius, K. Talavera, G. Owsianik, J. Prenen, G. Droogmans, and T. Voets, Gating of TRP channels: a voltage connection? J. Physiol. (2005). C.A. Perez, L. Huang, M. Rong, J.A. Kozak, A.K. Preuss, H. Zhang, M. Max, and R.F. Margolskee, A transient receptor potential channel expressed in taste receptor cells, Nat. Neurosci. 5 (2002) 1169–1176. D. Prawitt, T. Enklaar, G. Klemm, B. Gartner, C. Spangenberg, A. Winterpacht, M. Higgins, J. Pelletier, and B. Zabel, Identiﬁcation and characterization of MTR1, a novel gene with homology to melastatin (MLSN1) and the trp gene family located in the BWS-WT2 critical region on chromosome 11p15.5 and showing allele-speciﬁc expression, Hum. Mol. Genet. 9 (2000) 203–216. D. Prawitt, M.K. Monteilh-Zoller, L. Brixel, C. Spangenberg, B. Zabel, A. Fleig, and R. Penner, TRPM5 is a transient Ca2+-activated cation channel responding to rapid changes in [Ca2+]i, Proc. Natl. Acad. Sci. USA 100 (2003) 15166–15171. T. Rohacs, C.M. Lopes, I. Michailidis, and D.E. Logothetis, PI(4,5)P(2) regulates the activation and desensitization of TRPM8 channels through the TRP domain, Nat. Neurosci. 8 (2005) 626–634. K. Talavera, K. Yasumatsu, T. Voets, G. Droogmans, N. Shigemura, Y. Ninomiya, R.F. Margolskee, and B. Nilius, Heat activation of TRPM5 underlies thermal sensitivity of sweet taste, Nature 438 (2005) 1022–1025. N.D. Ullrich, T. Voets, J. Prenen, R. Vennekens, K. Talavera, G. Droogmans, and B. Nilius, Comparison of functional properties of the Ca2+-activated cation channels TRPM4 and TRPM5 from mice, Cell Calcium 37 (2005) 267–278. P. Varnai and T. Balla, Visualization of phosphoinositides that bind pleckstrin homology domains: calcium- and agonist-induced dynamic changes and relationship to myo-[3H]inositol-labeled phosphoinositide pools, J. Cell Biol. 143 (1998) 501–510. T. Voets, G. Droogmans, U. Wissenbach, A. Janssens, V. Flockerzi, and B. Nilius, The principle of temperature-dependent gating in cold- and heat-sensitive TRP channels, Nature 430 (2004) 748–754. X.Z. Xu, F. Moebius, D.L. Gill, and C. Montell, Regulation of melastatin, a TRPrelated protein, through interaction with a cytoplasmic isoform, Proc. Natl. Acad. Sci. USA 98 (2001) 10692–10697. G. Yellen, Single Ca2+-activated nonselective cation channels in neuroblastoma, Nature 296 (1982) 357–359. Y. Zhang, M.A. Hoon, J. Chandrashekar, K.L. Mueller, B. Cook, D. Wu, C.S. Zuker, and N.J. Ryba, Coding of sweet, bitter, and umami tastes: different receptor cells sharing similar signaling pathways, Cell 112 (2003) 293–301. Z. Zhang, H. Okawa, Y. Wang, and E.R. Liman, Phosphatidylinositol 4,5-bisphosphate rescues TRPM4 channels from desensitization, J. Biol. Chem. 280 (2005) 39185–39192. B. Nilius, F. Mahieu, J. Prenen, A. Janssens, G. Owsianik, R. Vennekens, and T. Voets, The Ca2+-activated cation channel TRPM4 is regulated by phosphatidylinositol 4,5-biphosphate, Embo. J. 25 (2006) 467–478.

16

Genetics Can Be Painless: Molecular Genetic Analysis of Nociception in Drosophila W. Daniel Tracey, Jr. Duke University Medical Center

CONTENTS A Drosophila Model for Nociception ...................................................................214 Mutations in a TRPA Ion Channel Increase Mechanical and Thermal Nociception Thresholds ...................................................................214 Distinct Cellular and Molecular Pathways for Innocuous and Noxious Touch Detection in Drosophila .......................................................216 Thermo TRP Channels Are Temperature Sensors in Mammals...........................217 Multiple TRPA Channels Are Heat Activated in Flies .........................................218 Evidence for Combinatorial Encoding of Temperature Response .......................218 Putative Functions for the High-Threshold and Low-Threshold Types of Thermosensory Neurons ...................................................................................221 The Mammalian TRPA1 Channel Is Activated by Irritants That Elicit Sensations of Burning Pain.................................................................222 TRP Channels Have Been Implicated in Mechanotransduction ..........................222 References..............................................................................................................223

Pain is a major health problem. Despite the importance of adequate pain modulation in relieving patients’ suffering, the available pharmacological agents for treating pain are limited. Existing pain-treating drugs that predominantly target the cyclooxygenase and opioid pathways are often inadequate in providing relief from pain and can have undesirable side effects. A ﬁrst step in improving the armamentarium of physicians in treating pain is identifying new molecular targets for analgesic drugs. In this chapter, I describe a system for using Drosophila as a platform for discovering genes required for the function of insect nociceptive sensory neurons.1 Forward genetics using Drosophila has resulted in important mechanistic insights into a variety of nervous system processes for which suitable behavioral 213

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assays have been developed. While these contributions are too numerous to list, notable examples include identiﬁcation of the period locus affecting circadian rhythms, which led to a detailed understanding of molecular clocks,2 and mutations at the shaker locus that led to the cloning of the ﬁrst voltage-activated K+ channel.3

A DROSOPHILA MODEL FOR NOCICEPTION For studies of nociception, my colleagues and I developed a paradigm based on a behavioral output that occurs in Drosophila larvae that have been stimulated with noxious heat or noxious mechanical stimuli. In response to such stimuli, the larvae produce a stereotyped escape response.1 In contrast to normal locomotion, where the larva moves with caudal to rostral peristaltic waves of muscle contraction, after a noxious heat or mechanical stimulus larval locomotion changes to a characteristic “writhing” pattern. The defensive pattern of locomotion causes the larvae to roll across the substrate.1 The threshold probe temperature for eliciting the Drosophila nociception behavior is 39°C. Similarly, through extracellular electrophysiological recordings we identiﬁed temperature-sensitive neurons of the peripheral nervous system that initiate ﬁring at approximately 39°C.1 Importantly, 39°C is similar to the threshold temperature of mammalian nociceptive nerve endings.4 The similarity in temperature thresholds in mammalian nociceptive nerve endings and in insect nociception may indicate that nociception evolved in an ancient common ancestor of insects and mammals to protect a vital cellular process that begins to fail at or above approximately 39°C. Our goal is to identify precisely the signaling mechanisms that evolved to acutely protect the organism from this critical failure. While I was a postdoc in the laboratory of Seymour Benzer, I took advantage of the stereotyped larval nociception behavioral response to conduct a genetic screen for mutations that produce a delayed behavioral response to noxious heat. In this screen, approximately 50 lines with altered nociception were identiﬁed.

MUTATIONS IN A TRPA ION CHANNEL INCREASE MECHANICAL AND THERMAL NOCICEPTION THRESHOLDS The mutant we characterized in greatest detail was named painless.1 We found that larvae mutant for painless have an increased threshold for eliciting the defensive behavioral response to either noxious heat or strong mechanical stimuli. Approximately 90 percent of wild-type larvae responded to a 46°C stimulus in 0.4 second.1 In contrast, the average response latency is greater than three seconds in painless mutant larvae. The delayed behavioral response of painless mutant larvae is not due to a defective motor output because the mutant larvae show a rapid (0.4 second) response to a hotter (53°C) probe. 1 In wild-type larvae, the defensive rolling behavior also occurred in response to a 30mN punctate mechanical stimulus. In contrast, an equivalent level of response in painless mutant animals required a 100mN stimulus.1 These results indicate that the painless mutation results in increased thresholds for both thermal and mechanical nociception.

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In electrophysiological recordings, we found that bulk spiking activity increased three-fold in the noxious temperature range (>39°C) in wild-type larval nerves and that this increase of spiking did not occur in the painless mutant larvae. In wild type, at least two classes of neurons responded to increases in temperature. One class (low threshold) showed spontaneous room temperature ﬁring and a marked increase in ﬁring from 29–39°C. A second class (high threshold) was silent at room temperature and initiated strong ﬁring above 39°C. Both classes of temperature response were absent in recordings from painless mutant nerves.1 We rescued the painless mutant phenotype with 8.5Kb of genomic DNA containing a single predicted open reading frame (ORF). By in situ hybridization using cDNA probes encoding the ORF, we found that the transcript was expressed in the larval multidendritic neurons. In addition, an antibody raised against a peptide within the painless protein detected punctate structures associated with the dendritic arbor of multidendritic neurons. We isolated an enhancer trap allele in which the yeast transcription factor GAL4 was inserted at the painless locus. We used this GAL4 line to drive expression of green ﬂuorescent protein (GFP) under control of GAL4-binding sites (UAS sites) and again observed expression in both the multidendritic neurons and chordotonal neurons. Interestingly, multidendritic neurons of Drosophila larvae have morphological similarity to naked nerve endings of mammalian nociceptors in that they are apparently not associated with a specialized receptor cell and have neurites that are in direct contact with epidermal cells that secrete cuticle (Figure 16.1).1

FIGURE 16.1 Type II (multidendritic) sensory neurons. While the function of these cells was previously unknown, the available evidence suggests that these cells, or a subset of them, function as nociceptors. Blocking the synaptic output of these cells blocks nociception behavior; these cells are activated by heat and express the painless gene. In contrast, genes required for light touch detection are not expressed in the multidendritic neurons.

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Several lines of evidence suggest that the multidendritic neurons, but not the chordotonal neurons, included the heat-sensing nociceptive neurons. First, blocking the synaptic output of the multidendritic neurons with tetanus toxin completely abrogates the behavioral response to noxious heat.1 Second, multidendritic neurons show strong Ca++ responses to heating with only weak responses in chordotonal neurons.5 Third, atonal mutants, which lack chordotonal neurons, showed a rapid response to noxious heat.1 The painless gene encodes an ion channel of the transient receptor potential (TRP) family.1 Isolation of mutations in a TRP ion channel in our screen is consistent with other ﬁndings that have implicated individual TRP channels in mechanotransduction1,6–10 and in thermal sensory transduction.1,11–16 Based on sequence homology, the TRP channel family has seven subfamilies: TRPA, TRPV, TRPP, TRPM, TRPC, TRPN, and TRPML.17 The painless gene is a member of the TRPA subfamily.

DISTINCT CELLULAR AND MOLECULAR PATHWAYS FOR INNOCUOUS AND NOXIOUS TOUCH DETECTION IN DROSOPHILA When we initiated our nociception screens, Kernan and colleagues had previously screened for chemically induced mutations that resulted in larval insensitivity to a light touch.18 In this paradigm, the larvae are lightly touched on the head segment using an eyelash that elicits a maximum force of <10mN. In response, wild-type larvae pause forward movement or initiate reverse peristaltic waves of muscle contraction to move away from the stimulus. At the behavioral level, this response is completely distinct from the writhing nociception response that occurs with harsh touch of >30mN. Kernan and others have cloned several light touch–insensitive mutants such as NompC, NompA, and unc.6,19,20 Each of these genes has been expressed in ciliated Type I bipolar sensory neurons or in other cells of the Type I sensillum (Figure 16.2). Touch-insensitive mutants isolated in the Kernan screen do not affect the writhing response to strong mechanical stimuli (W.D. Tracey, unpublished). In contrast, the painless gene product is primarily absent from Type I sensory neurons but is strongly expressed in the multidendritic Type II sensory neurons. Consistent with this, painless mutants show a normal behavioral response to light touch but are defective in the strong touch response. This result genetically separates the light touch response from the harsh touch response. Because light touch response genes are absent from the Type II sensory neurons while painless is present in these cells, these data further implicate the Type II sensory neurons as nociceptors. Taken together, the available evidence suggests that Drosophila Type I mechanosensory neurons are required for light touch detection, while Type II sensory neurons are used in harsh touch detection.

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FIGURE 16.2 Type I sensory neurons. These bipolar sensory neurons function as part of a sensillum consisting of four cells. Type I sensory neurons are used in a variety of sensory processes including olfaction, gustation, light touch detection, and in hearing of adult ﬂies. Shown here are ﬁve chordotonal neurons used in the larva as stretch receptors and in the adult for hearing. These cells were detected by a green ﬂuorescent protein reporter.

THERMO TRP CHANNELS ARE TEMPERATURE SENSORS IN MAMMALS In mammals, external temperature is sensed by the nerve endings of neurons whose cell bodies reside in sensory ganglia such as the trigeminal and dorsal root ganglia (DRG). Several TRP channels have been expressed in trigeminal and DRG neurons and have been shown to produce temperature-dependent currents in heterologous expression systems. Indeed, the ﬁrst identiﬁed temperature-sensitive TRP channel (TRPV1) is activated by temperatures >42°C. TRPV1 is also activated by capsaicin, the spicy ingredient in chili peppers.12 In addition, mice mutant for TRPV1 eat capsaicin with impunity. Combined, these observations indicate that the ‘‘hot’’ sensation produced by capsaicin and other vanilloid compounds is due to activation of thermal pain-sensing nerve endings through TRPV1.12 Other warmth-activated TRPs include TRPV2 (>52°C),11 TRPV3 (>31°C),21 and TRPV4 (>25°C).22 TRPM8 is activated by menthol and cooling (8–28°C).14

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According to the properties of these channels in heterologous expression systems, the best candidate for encoding the temperature-mediated response of nociceptive sensory neurons in mice is TRPV1. This is because heat-activated nociceptors show increased ﬁring at temperature thresholds when the nerve endings reach a temperature of approximately 39°C, and the activation threshold for TRPV1 is similar. Indeed, thermal hyperalgesia is signiﬁcantly reduced in TRPV1 knockout mice, but thermal nociception is not completely abrogated by the TRPV1 mutation.23,24 More recently, mice mutant for TRPV3 have also been shown to have altered thermal preference and increased nociception thresholds.25

MULTIPLE TRPA CHANNELS ARE HEAT ACTIVATED IN FLIES Drosophila TRPV channels have not been found to be heat activated.8,26 In contrast, two of the three identiﬁed TRPA channels in ﬂies are activated by heat.27–29 Phylogenetic analysis suggests that more than one TRPA gene existed in a common ancestor of ﬂies and mammals. The presence of a single TRPA gene in modern humans, rats, and mice suggests either a simpliﬁcation of this gene family in the mammalian lineage or an expansion of the family in the insect lineage. Three TRPA genes in ﬂies have been functionally analyzed and have apparently distinct functions. As mentioned above, painless is required for thermal nociception but not for thermotaxis.29,30 The painless gene that we identiﬁed by forward genetics is not the orthologue of the mammalian TRPA1 channel. Another gene, dTRPA1, is more closely related to mammalian TRPA1.29,30 RNAi knockdown of dTRPA1 results in thermotaxis defects without affecting thermal nociception.29 Mutants of a third Drosophila TRPA gene named pyrexia show reduced tolerance to thermal stress.28 Although painless is not the Drosophila sequence orthologue of mammalian TRPA1, the expansion of the TRPA family found in ﬂies may have allowed each Drosophila TRPA gene to retain a subfunction of an ancestral TRPA gene. Because painless is expressed in Drosophila nociceptors but dTRPA1 is not, painless can be considered the functional orthologue of mammalian TRPA1 when nociception pathways are under consideration. It is possible that pyrexia may also be involved in nociception because it is coexpressed in multidendritic neurons with painless, but we have not observed nociception defects in pyrexia mutants (W.D. Tracey, unpublished).

EVIDENCE FOR COMBINATORIAL ENCODING OF TEMPERATURE RESPONSE The painless channel is required for Drosophila sensory neurons to be activated by increasing temperature.1 However, paradoxically, painless is required in vivo for the heat-dependent activation of two distinct types of thermosensory neurons (Figure 16.3).1 The ﬁrst type (low threshold) activates at approximately 29°C and inactivates at around 39°C (Figure 16.3). The second type (high threshold) activates at 39°C but is silent below this temperature. Several distinct hypotheses can be proposed to explain the ﬁring properties of these two classes of thermosensory neurons. The ﬁrst possibility is that there are multiple isoforms of painless that show distinct activation

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FIGURE 16.3 Schematic representation of Drosophila thermosensory neuron ﬁring patterns. A low-threshold type has spontaneous room temperature activity and increased ﬁring above 29°C and inactivation at 39°C. A high-threshold type activates at approximately 39°C. Temperature-activated ﬁring of both the low-threshold type and high-threshold type of sensory neuron is absent in painless mutant nerves. In wild-type, thermosensory neurons have the largest amplitude spikes, with spontaneous ﬁring at room temperature, and are detectable in painless mutant nerves (bottom trace) but are not activated by high temperature.

thresholds of 29°C and 39°C, respectively. Evidence from expressed sequence tags supports the possibility that there may be multiple painless isoforms. Our preliminary unpublished results suggest that at least one of the painless isoforms, when expressed heterologously, is directly heat activated and has an activation threshold of 29°C. In addition, we identiﬁed a second isoform of painless that is a candidate for encoding the putative 39°C threshold isoform. However, the pyrexia thermoTRP also has similar multiple isoforms, and thermal-activation thresholds among these isoforms is similar,28 so it may be unlikely that the two distinct isoforms of painless have dramatically different thermal-activation thresholds. A second model that would explain the properties of these two types of thermosensory neurons is more likely (Figure 16.4). This model invokes other thermosensory

FIGURE 16.4 Hypothesis for combinatorial encoding of thermosensory neuron ﬁring properties. We propose that painless is an “excitatory” thermoTRP and that pyrexia is an inhibitory thermoTRP. This model predicts that the low-threshold type of thermosensory neuron coexpresses painless and pyrexia. Further, we propose that the high-threshold type of thermosensory neurons results from coexpression of painless and another thermosensory K channel (such as dTRPA1).

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channels. In this model, there is only a single thermosensitive painless channel, which is activated at temperatures >29°C, and this painless channel is expressed in both the low-threshold and high-threshold types of thermosensory neurons. The properties of the high-threshold cell could be explained if there was a coexpressed K+ current in that type of cell that would prevent ﬁring below 39°C. An interesting candidate for such a channel could be a leak channel that showed K+ selectivity and inactivation at >39°C. A second K+-selective thermoTRP channel that opens at >39°C could explain the inactivation of the low-threshold thermosensory neuron at this temperature. A variation of this model is that painless may form heteromultimers with these other channels and the heteromultimer might show a thermal-activation threshold distinct from that of the painless homomultimer. Both dTRPA1 and pyrexia have been shown to function as thermoTRPs and are thus candidates as these hypothetical K+-selective thermoTRPs or as heteromeric partners for painless. dTRPA1 is activated at temperatures greater than 23°C, while pyrexia is activated at temperatures greater than 39°C. Indeed, pyrexia-PA and pyrexia-PB heteromeric channels are somewhat selective for K+ and have been proposed to inhibit neurons from ﬁring under conditions of thermal stress.28 The expression pattern, thermal-activation threshold, and ion selectivity of pyrexia are consistent with the possibility that pyrexia is a channel that is coexpressed with painless to inactivate the low-threshold type of thermosensory neuron at 39°C. This is because pyrexia is expressed in a subset of the multidendritic neurons that also express painless.28 A logical extension of this model is that dTRPA1 could be the other putative K+-selective thermoTRP. However, currently available anti-dTRPA1 antibodies have not detected expression in multidendritic neurons. This may be due to levels of expression below the limits of detection or, alternatively, other K+selective channels must be considered. A novel and exciting prediction of the second model is that the ﬁring properties of Drosophila thermosensitive neurons might be combinatorially determined by “excitatory” and “inhibitory” thermoTRPs. Based on the thermal thresholds of the known Drosophila thermoTRPs and on the painless mutant phenotypes, we hypothesize that painless is an excitatory TRPA channel and that pyrexia and possibly dTRPA are inhibitory thermoTRPs. A simple combinatorial code could explain the ﬁring properties of the neurons. Namely, coexpression of painless and pyrexia might encode the lowthreshold ﬁring pattern and coexpression of dTRPA1 (or another thermo K+ channel), and painless might encode the high-threshold ﬁring pattern (Figure 16.4). This is a theoretical prediction, and it is possible that the inhibitory K+-selective channels may not be TRP channels at all. For example, the seizure gene encodes a K+ channel that when mutant causes neuronal hyperexcitability in ﬂies, speciﬁcally at high temperatures. The simplicity of the Drosophila peripheral nervous system provides a signiﬁcant advantage to understand neural substrates encoding thermosensory information. Most important, the precise number, identity, and location of individual peripheral neurons are invariant from animal to animal. This allows us to determine the precise expression patterns of painless, dTRPA1, and pyrexia to the resolution of individual identiﬁed neurons. Further, using the tools available in Drosophila, we will be able to determine the consequence of altering the activity of these identiﬁed neurons on thermal preference and nociception behaviors.

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PUTATIVE FUNCTIONS FOR THE HIGH-THRESHOLD AND LOW-THRESHOLD TYPES OF THERMOSENSORY NEURONS The two types of thermosensory neurons that we discovered in Drosophila suggest a mechanism for the brain to compute the direction of temperature change during a noxious thermal stimulus presentation (Figure 16.5). For example, central neurons receiving input from both the low-threshold type and the high-threshold type could compute that temperature is increasing in instances where ﬁring of the lowthreshold type temporally precedes the high-threshold type. The reverse pattern would indicate a temperature decrease. Although intuitively obvious, whether the brain of the ﬂy or any other species actually uses this simple mechanism of comparison is not known. This hypothesis for computation of direction of temperature change makes interesting predictions concerning the effects of removing the low-threshold neuron. For example, an animal lacking a signal from this neuron may show a prolonged rolling response to a noxious heat probe. This would occur because the high-threshold neuron response should be sufﬁcient to initiate the rolling behavior, but the brain may require input from the low-threshold neuron in order to “know” when the thermal danger has passed and to reinitiate normal locomotion. Absence of input from the low-threshold neuron would therefore be predicted to cause a form of hyperalgesia. That human central nervous system pain pathways involve comparison of input from distinct thermosensory neuronal types is suggested from the thermal grill pain illusion. In this illusion, a human hand placed on a grill of alternating cool (18°C) and warm (37°C) bars elicits a sensation of pain.31–33 This suggests that innocuous thermoreception inﬂuences central pain circuits.

FIGURE 16.5 Hypothetical thermal nociception circuit. We hypothesize that the lowthreshold type of thermosensory neuron (top trace) drives inhibition of a rolling locomotion pattern generator. The high-threshold type (bottom trace) should simultaneously activate rolling locomotion and drive inhibition of normal locomotion.

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THE MAMMALIAN TRPA1 CHANNEL IS ACTIVATED BY IRRITANTS THAT ELICIT SENSATIONS OF BURNING PAIN As we characterized the painless mutant, other groups of investigators simultaneously began to investigate the single homologue of the painless gene that has been identiﬁed in mammalian genomes (TRPA1).34 The available evidence suggests that the TRPA1 gene is in fact an excellent candidate to play an important role in pain signaling. First, TRPA1 was found to be expressed in a subset of TRPV1-expressing neurons, which are likely to be nociceptors.34 Second, using heterologous expression systems, the TRPA1 gene was found to be activated by a variety of compounds known to elicit a burning sensation of pain such as wasabi (Japanese horseradish), mustard oils, cinnemaldehyde (the active compound of cinnamon and spicy hot candies), and, most recently, raw garlic.35,36,37,46 Activation of TRPA1 in vitro by these compounds suggests that the psychophysical sensation produced by activating TRPA1-expressing neurons in vivo is that of burning pain. In addition, TRPA1 can be activated downstream of pain-modulating G-protein-coupled receptors such as bradykinin receptors.35,36 mTRPA1 may function as a thermoTRP. The mouse channel can be activated by noxious cold (<16°C) in heterologous expression systems,38 but this has not been observed in every study of this channel.35,36 Whether noxious cold activation of mTRPA1 is direct or due to an indirect effect of elevated intracellular Ca++ is not known.35,36 In culture, noxious cold–responding neurons do not respond to mustard oils, which do activate TRPA1 in heterologous cell types.39 The observation that compounds that elicit a burning hot sensation activate TRPA1 in heterologous cells may indicate that in vivo functions of TRPA1 are present in neurons that participate in noxious heat detection, or, alternatively, burning cold and burning heat may be psychophysically related cognitive processes.38

TRP CHANNELS HAVE BEEN IMPLICATED IN MECHANOTRANSDUCTION The TRP channels osm-9, NOMP-C, Nanchung, Inactive, TRPV1, zNOMP-C, and painless are each genetically required for behavioral or physiological responses to mechanical stimuli.1,6–9,26 In C. elegans, osm-9 mutants are defective in nose touch avoidance and for other nociceptive functions such as avoidance of high osmotic strength.7 In Drosophila, no mechanoreceptor potential-C mutants are defective in the response to touch and for the mechanoreceptor potential of neurons that innervate bristles of the ﬂy.6,18 Drosophila Nanchung and Inactive, and zebraﬁsh NOMP-C, are each required for hearing.40–43 Finally, painless mutant larvae have an increased threshold for avoiding harsh touch but a normal response to light touch.1 Perhaps the longest-sought mechanotransducing ion channel is the channel responsible for depolarization of the hair cells of the inner ear in response to mechanical vibrations that are produced by sound. This channel is present in only several hundred copies per cell and thus has resisted biochemical puriﬁcation.

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Recent evidence suggests that, in addition to nociceptive sensory neurons, TRPA1 is present in hair cells of the inner ear. Antibodies raised against TRPA1 detect structures at the tips of actin-rich stereovilli in several species and knockdown reduced the receptor potential.10 Thus, TRPA1 is an excellent candidate for the hair cell mechanotransducer.10 This emerging evidence suggests that TRPA1 may have distinct functions in distinct cell types. In nociceptors, TRPA1 may elicit burning sensations of pain, and in the ear it is a good candidate to be involved in hair cell mechanotransduction. The function of TRPA1 in nociceptors may be as a “painful pinch” detector as with painless. The exact role of TRPA1 in these cell types awaits the generation of mice mutant in this gene. Nociceptive neurons and hair cells are two seemingly very distinct cell types that express TRPA1. It is likely that cell-type speciﬁc cofactors modulate the function of this channel in these distinct cells. Our Drosophila system is speciﬁcally designed to identify peripheral nervous system regulators of nociception. The Drosophila genome contains neither opioid receptors nor cyclooxygenase enzymes. While these pathways are clearly extremely important pathways in pain modulation, it is essential to identify novel pathways that could be modulated independently of them. Because Drosophila lacks these pathways, we have a priori knowledge that the genes we identify will not touch on the well-studied opioid and cyclooxygenase systems. The understanding of peripheral pathways for nociception and identiﬁcation of new targets that can modulate peripheral pathways is an important area of health research for improved patient care. Although many compounds used in pain management act on central nervous system targets, it is well established that blocking peripheral input to the dorsal horn can prevent wind-up (a sensitized neural circuit in the dorsal horn).44 Once windup has occurred, pain management for the physician becomes more difﬁcult.45 Therefore, understanding primary nociception is important to prevent sensitized neural circuits that may occur as a consequence of tissue damage.45 If the peripheral pathways can be modulated, this may prevent wind-up and the development of central pain syndromes.44,45 Physicians working on the incredibly complex problem of human pain in many instances simply do not have the pharmacological tools that they need for adequate patient care. By focusing on the simplest model system possible, we hope to contribute to a mechanistic understanding that will assist in providing these tools.

REFERENCES 1. Tracey, W.D., Jr. et al., Painless, a Drosophila gene essential for nociception. Cell, 2003. 113(2): 261–73. 2. Konopka, R.J. and S. Benzer, Clock mutants of Drosophila melanogaster. Proc. Natl. Acad. Sci. USA, 1971. 68(9): 2112–16. 3. Papazian, D.M. et al., Cloning of genomic and complementary DNA from Shaker, a putative potassium channel gene from Drosophila. Science, 1987. 237(4816): 749–53. 4. Tillman, D.B. et al., Response of C ﬁbre nociceptors in the anaesthetized monkey to heat stimuli: correlation with pain threshold in humans. J. Physiol., 1995. 485 (Pt. 3): 767–74.

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5. Liu, L. et al., Identiﬁcation and function of thermosensory neurons in Drosophila larvae. Nat. Neurosci., 2003. 6(3): 267–73. 6. Walker, R.G., A.T. Willingham, and C.S. Zuker, A Drosophila mechanosensory transduction channel. Science, 2000. 287(5461): 2229–34. 7. Colbert, H.A., T.L. Smith, and C.I. Bargmann, OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans. J. Neurosci., 1997. 17(21): 8259–69. 8. Kim, J. et al., A TRPV family ion channel required for hearing in Drosophila. Nature, 2003. 424(6944): 81–84. 9. Sidi, S., R.W. Friedrich, and T. Nicolson, NompC TRP channel required for vertebrate sensory hair cell mechanotransduction. Science, 2003. 301(5629): 96–99. 10. Corey, D.P. et al., TRPA1 is a candidate for the mechanosensitive transduction channel of vertebrate hair cells. Nature, 2004. 432(7018): 723–30. 11. Caterina, M.J. et al., A capsaicin-receptor homologue with a high threshold for noxious heat. Nature, 1999. 398(6726): 436–41. 12. Caterina, M.J. et al., The capsaicin receptor: a heat-activated ion channel in the pain pathway. Nature, 1997. 389(6653): 816–24. 13. Bassilana, F. et al., The acid-sensitive ionic channel subunit ASIC and the mammalian degenerin MDEG form a heteromultimeric H+-gated Na+ channel with novel properties. J. Biol. Chem., 1997. 272(46): 28819–22. 14. McKemy, D.D., W.M. Neuhausser, and D. Julius, Identiﬁcation of a cold receptor reveals a general role for TRP channels in thermosensation. Nature, 2002. 416 (6876): 52–58. 15. Peier, A.M. et al., A TRP channel that senses cold stimuli and menthol. Cell, 2002. 108(5): 705–15. 16. Jordt, S.E., D.D. McKemy, and D. Julius, Lessons from peppers and peppermint: the molecular logic of thermosensation. Curr. Opin. Neurobiol., 2003. 13(4): 487–92. 17. Clapham, D.E., TRP channels as cellular sensors. Nature, 2003. 426(6966): 517–24. 18. Kernan, M., D. Cowan, and C. Zuker, Genetic dissection of mechanosensory transduction: mechanoreception-defective mutations of Drosophila. Neuron, 1994. 12(6): 1195–206. 19. Baker, J.D., S. Adhikarakunnathu, and M.J. Kernan, Mechanosensory-defective, malesterile unc mutants identify a novel basal body protein required for ciliogenesis in Drosophila. Development, 2004. 131(14): 3411–22. 20. Chung, Y.D. et al., nompA encodes a PNS-speciﬁc, ZP domain protein required to connect mechanosensory dendrites to sensory structures. Neuron, 2001. 29(2): 415–28. 21. Smith, G.D. et al., TRPV3 is a temperature-sensitive vanilloid receptor-like protein. Nature, 2002. 418(6894): 186–90. 22. Guler, A.D. et al., Heat-evoked activation of the ion channel, TRPV4. J. Neurosci., 2002. 22(15): 6408–14. 23. Caterina, M.J. et al., Impaired nociception and pain sensation in mice lacking the capsaicin receptor. Science, 2000. 288(5464): 306–13. 24. Davis, J.B. et al., Vanilloid receptor-1 is essential for inﬂammatory thermal hyperalgesia. Nature, 2000. 405(6783): 183–87. 25. Moqrich, A. et al., Impaired thermosensation in mice lacking TRPV3, a heat and camphor sensor in the skin. Science, 2005. 307(5714): 1468–72. 26. Gong, Z. et al., Two interdependent TRPV channel subunits, inactive and Nanchung, mediate hearing in Drosophila. J. Neurosci., 2004. 24(41): 9059–66. 27. Viswanath, V. et al., Opposite thermosensor in fruitﬂy and mouse. Nature, 2003. 423(6942): 822–23.

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28. Lee, Y. et al., Pyrexia is a new thermal transient receptor potential channel endowing tolerance to high temperatures in Drosophila melanogaster. Nat. Genet., 2005. 37(3): 305–10. 29. Rosenzweig, M. et al., The Drosophila ortholog of vertebrate TRPA1 regulates thermotaxis. Genes Dev., 2005. 19(4): 419–24. 30. Tracey, W.D., Jr. et al., Quantitative analysis of gene function in the Drosophila embryo. Genetics, 2000. 154(1): 273–84. 31. Bouhassira, D. et al., Investigation of the paradoxical painful sensation (‘‘illusion of pain’’) produced by a thermal grill. Pain, 2005. 114(1-2): 160–67. 32. Craig, A.D. et al., Functional imaging of an illusion of pain. Nature, 1996. 384(6606): 258–60. 33. Craig, A.D. and M.C. Bushnell, The thermal grill illusion: unmasking the burn of cold pain. Science, 1994. 265(5169): 252–55. 34. Story, G.M. et al., ANKTM1, a TRP-like channel expressed in nociceptive neurons, is activated by cold temperatures. Cell, 2003. 112(6): 819–29. 35. Bandell, M. et al., Noxious cold ion channel TRPA1 is activated by pungent compounds and bradykinin. Neuron, 2004. 41(6): 849–57. 36. Jordt, S.E. et al., Mustard oils and cannabinoids excite sensory nerve ﬁbres through the TRP channel ANKTM1. Nature, 2004. 427(6971): 260–65. 37. Macpherson, L.J. et al., The pungency of garlic: activation of TRPA1 and TRPV1 in response to allicin. Curr. Biol., 2005. 15(10): 929–34. 38. Darboux, I. et al., A new member of the amiloride-sensitive sodium channel family in Drosophila melanogaster peripheral nervous system. Biochem. Biophys. Res. Commun., 1998. 246(1): 210–16. 39. Babes, A., D. Zorzon, and G. Reid, Two populations of cold-sensitive neurons in rat dorsal root ganglia and their modulation by nerve growth factor. Eur. J. Neurosci., 2004. 20(9): 2276–82. 40. Mogil, J.S. et al., One or two genetic loci mediate high opiate analgesia in selectively bred mice. Pain, 1995. 60(2): 125–35. 41. Marek, P., I. Panocka, and B. Sadowski, Selective breeding of mice for high and low swim analgesia: differential effect on discrete forms of footshock analgesia. Pain, 1987. 29(3): 393–98. 42. Bier, E. et al., Searching for pattern and mutation in the Drosophila genome with a P- lacZ vector. Genes Dev., 1989. 3(9): 1273–87. 43. Pastor, J., B. Soria, and C. Belmonte, Properties of the nociceptive neurons of the leech segmental ganglion. J. Neurophysiol., 1996. 75(6): 2268–79. 44. Seltzer, Z. et al., The role of injury discharge in the induction of neuropathic pain behavior in rats. Pain, 1991. 46(3): 327–36. 45. Arendt-Nielsen, L. and S. Petersen-Felix, Wind-up and neuroplasticity: is there a correlation to clinical pain? Eur. J. Anaesthesiol. Suppl., 1995. 10: 1–7.

17

TRPV Family Ion Channels and Other Molecular Components Required for Hearing and Proprioception in Drosophila Changsoo Kim Chonnam National University

CONTENTS Abstract ..................................................................................................................227 Introduction............................................................................................................228 The Drosophila Chordotonal Organs ....................................................................229 Antennal and Nonantennal Chordotonal Organs ......................................229 Scolopidium Structure ...............................................................................230 Key Gene Products Involved in Mechanosensation.................................. 231 Gene Products Required for Both Auditory and External Sensory Transduction.......................................................................231 Gene Products Involved in Auditory Transduction, but Not in External Sensory Transduction ......................................233 Drosophila TRPV Channels: Nanchung and Inactive ..........................................234 Nanchung and Inactive ..............................................................................234 How Is Nan-Iav Gated in Vivo?................................................................235 Concluding Remarks .............................................................................................237 Acknowledgments..................................................................................................237 References..............................................................................................................237

ABSTRACT In Drosophila, chordotonal sensory organs mediate hearing and proprioception (the body’s awareness of motion and spatial orientation). However, little is known about the mechanisms by which these organs convert mechanical force into the two aforementioned senses. Sensory cilia on the chordotonal neurons are mechanosensitive

227

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organelles that carry out mechanical transduction. Recently, various molecular components required for cilia formation and function have been identiﬁed. Moreover, two subunits of the Drosophila TRPV (vanilloid receptor–related transient receptor potential) family of ion channels were localized to the sensory cilia of the chordotonal neurons. Current evidence suggests that Drosophila TRPV channels form a mechanotransducer complex that converts mechanical force into a neuronal electrical signal.

INTRODUCTION Mechanosensation involves the transduction of mechanical stimuli into neuronal impulses (mechanotransduction) and is necessary for hearing, touch, and proprioception (the body’s awareness of limb position and movement). Mechanosensation occurs in sensory neuron–associated mechanosensitive organelles, such as the cilia in Drosophila chordotonal sensory organs and the stereocilia (hair bundles) on auditory hair cells in the cochlea (the vertebrate organ involved in hearing). Mechanosensitive ion channels are presumed to be located in the mechanosensitive organelles to convert mechanical force into receptor potential (that is, depolarization of the sensory neuron). Identiﬁcation of the molecular components of mechanosensitive organelles and channels is crucial if one is to decipher how mechanical force is transformed into the electrical activity of neurons. A number of the genes involved in cilia formation were identiﬁed recently in Drosophila,1,2 and candidate mechanosensitive channel components have been pinpointed in both invertebrates and vertebrates.3–8 The Drosophila chordotonal organs mediate hearing and proprioception and are evolutionarily and functionally related to vertebrate cochleae.2 Although the gross morphologies of chordotonal organs and cochleae differ, these related structures may be derived from a common ancestor (Figure 17.1). This notion is supported by the following observations. First, the receptor neurons of both organs have true cilia at their apical ends.8 Second, although the mechanosensitive organelle of the chordotonal organs—the sensory cilia—are tubulin based and the mechanosensitive organelle of cochleae—the stereocilia—are actin based, both structures are bathed in an endolymph with an unusual composition (high K+, low Ca2+).1,9 Third, the receptor neurons, which become depolarized and transmit electrical signals in response to mechanostimulation, are speciﬁed by the orthologous proneural genes atonal and Math1, in Drosophila and mammals, respectively.10–12 Drosophila atonal has been shown to specify auditory hair cells in mice,13 and mouse Math1 can substitute for atonal in forming the Drosophila chordotonal organs.13,14 Recently, candidate mechanical signal transducers were identiﬁed in Drosophila sensory cilia and vertebrate stereocilia and were shown to be members of the transient receptor potential (TRP) superfamily of ion channel subunits.15–17 In this chapter, I focus on the Drosophila chordotonal organs and their candidate transducer proteins Nanchung (Nan) and Inactive (Iav), which are members of the vanilloid receptor–related transient receptor potential (TRPV) subfamily.

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FIGURE 17.1 Schematic drawing of Drosophila and vertebrate auditory sensory cells. (A) Drosophila chordotonal neurons. The dark gray lines denote the location of Nan and Iav channel proteins. (B) A vertebrate hair cell. The dark gray marks the location of TRPA1.8 True cilia are shaded in light gray. The double arrows indicate the direction of force. DC, dendritic cap; CD, ciliary dilation; SR; scolopale rods; C, cilia; SC, stereocilia; TM, tectorial membrane; KC, kinocilium.

THE DROSOPHILA CHORDOTONAL ORGANS ANTENNAL

AND

NONANTENNAL CHORDOTONAL ORGANS

The chordotonal organs in Drosophila can be divided into two classes: nonantennal and antennal.2 Nonantennal chordotonal organs are widely distributed in larval cuticles—the hardened protective covering of many insects—and in the joints of adult appendages, and these organs function in vibration reception and proprioception. For example, the proximal femoral chordotonal organs (FCO) extend for the entire length of the femur and are apically attached to the femoro-tibial joint. This positioning allows the organs to sense both bending and extension of the tibia in the leg of the ﬂy.18,19 Drosophila antennal chordotonal organs (also called Johnston’s organs, JOs) are housed in the second segments of the antennae and are responsible for hearing (Figure 17.2).12,20 JOs span the second segment and apically attach to the joints of the second and third segments, thereby allowing the detection of third-segment movements. The third segment of the antenna is bonded to a featherlike structure known as the arista, which constitutes the sound receiver.21,22 The arista and third antenna segment vibrate in response to the “love song” elicited by courting males,

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FIGURE 17.2 The Drosophila ear. (A) A scanning electron microscopy picture of a Drosophila head (left). The segments of the antenna are numbered. A transmission electron microscopy picture of the second segment (right). A cuticle from the third segment is indicated with a star. The third segment vibrates when sound is introduced, and this movement gives rise to force, which is inﬂicted on the chordotonal cilia, as indicated by the double-headed arrow. (B) Transverse sections of scolopidia from the proximal to the apical region. C, cilia; SR, scolopale rods; CD, ciliary dilation; DC, dendritic cap.

which has an amplitude of 90 dB and a frequency of 160 Hz.23–26 The vibrations of the sound receiver stretch or bend the chordotonal cilia, thereby directly activating this mechanosensory organelle.1,4

SCOLOPIDIUM STRUCTURE Chordotonal organs are composed of hundreds of parallel copies of the scolopidium, a functional and structural unit that is unique to insects and crustaceans.27 A single scolopidium is composed of bipolar sensory receptor neurons and accessory cells arrayed in the following, very speciﬁc manner.1,28 The sensory neurons are surrounded by scolopale cells, which are attached to the basal and apical surfaces of cuticles by ligaments and attachment cells, respectively. The attachment and ligament cells are enriched in microtubules that are positioned in longitudinal arrays and thus are strongly stained with an antibody to α-tubulin, the main component of microtubules.29 These cells are, in turn, attached to two neighboring cuticles, which results in scolopidia that are linked to two separate movable cuticles.1 In this conﬁguration, chordotonal organs are able to sense the relative displacement of movable body parts.

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The bipolar sensory neuron bears an apically located dendrite from which a sensory cilium protrudes.27,30 The cilium extends into the surrounding endolymph and is attached at its distal tip to a dendritic cap (Figure 17.2). The dendritic cap— an extracellular matrix structure that consists of substances secreted by the scolopale cells—is attached to the attachment cells, which transmit mechanical force to the cilia via the dendritic cap. The cilia and endolymph are encased in a spindle-shaped cage made of prominent scolopale rods, which are intracellular structures that are deposited at the apical inner surface of scolopale cells. The scolopale rods are composed of longitudinal arrays of actin-rich materials with scattered microtubules. The ciliary axoneme—a ﬁber bundle that forms the central core of a cilium—displays a “9 + 0” microtubule arrangement, which is composed of nine longitudinal microtubule doublets and associated structures and lacks a central microtubule pair. As one proceeds longitudinally along a cilium, its diameter remains constant until one reaches the point of ciliary dilation,27,28 at which the regular array of the axoneme is broken by the existence of uncharacterized, electron-dense material on parallel microtubules.

KEY GENE PRODUCTS INVOLVED

IN

MECHANOSENSATION

Gene Products Required for Both Auditory and External Sensory Transduction Several molecules required for auditory transduction have been identiﬁed recently with forward and reverse genetics (described in detail below). Most of these molecules are required for the synthesis of cilia. In the absence of these molecules, cilia are malformed, split, or absent, and the sound-evoked potential, which measures the auditory function of chordotonal organs, is impaired, thus illustrating the importance of cilia in mechanotransduction.1,2 Because cilia are also important for the function of sensory touch bristles, which coat the ﬂy’s external surface, the absence of many of the same molecules leads to an impairment in the ﬂy’s ability to respond to touch sensations. Drosophila uncoordinated (unc) mutants have aberrant cilia, do not exhibit mechanotransduction, and produce nonmotile sperm.31 These observations imply that the Unc protein is required for ciliogenesis. Unc resides in two locations in the ﬂy: (1) in basal bodies, which lie at the base of and constitute templates for the synthesis of cilia and (2) in the paired centrioles of dividing spermatocytes.31 The Unc protein contains several short coiled-coil segments, proline-rich regions, and a Lissencephaly 1 homology (LisH) motif. In unc mutant ﬂies, the ciliary axonemes of chordotonal organs are missing, split, or truncated.31 The Drosophila pericentrin-like protein (D-PLP) is associated with centrioles and pericentriolar material (PCM).32 D-PLP is the only polypeptide in Drosophila that contains a pericentrin/AKAP450 centrosomal targeting (PACT) domain. D-PLP recruits most of the PCM components to the centrosome, a small halo of cytoplasm that surrounds the nucleus. The centrosome includes the centrioles and has a role in the organization of microtubules. In d-plp mutants, the PCM components are aberrantly recruited to mitotic spindle poles. In addition, the mechanosensory cilia

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are either absent or short and kinked, which underlies the lack of sound-evoked potential in these mutant ﬂies.32 The no-mechanoreceptor-potential A (nompA) gene is required for both bristle and chordotonal function.33 The NompA protein is expressed by the support cells that ensheath the sensory cilia and is localized to the extracellular dendritic cap in both the external sensory and chordotonal organs. NompA contains several plasminogen N-terminal (PAN) modules and a zona pellucida domain (ZP), each of which is found in protein components of the extracellular matrix.34–36 Tectorin, a component of the tectorial membrane of the cochlea, is a ZP domain–containing protein that overlies the cochlear hair cells and delivers mechanical stimuli to hair bundles.37 Mutations in α-tectorins result in detachment of the tectorial membrane from the hair cells and cause deafness in mice.37,38 Likewise, in nompA mutant ﬂies, the cilia are detached from the dendritic cap, which suggests that NompA functions in the crucial cilia-to-cap attachment.33 The no mechanoreceptor potential (nompB) gene encodes the Drosophila homologue of Chlamydomonas IFT88, Caenorhabditis elegans OSM-5, and mammalian Polaris/Tg737, all of which are members of “intraﬂagellar transport” (IFT) complexes.39 The IFT complex functions in cilia construction by transporting their components to the appropriate physiological locations. First described in the singlecelled alga Chlamydomonas, IFT is a process used to build and maintain ﬂagella and involves the bidirectional movement of uniformly sized particles along the ﬂagellum.40–42 Particle movement toward the tips of cilia and ﬂagella uses the kinesin II protein, and backward locomotion toward the cell body employs nonaxonemal dynein.43 The NOMPB protein is localized to sensory cilia and basal bodies. Mutations in nompB give rise to either malformed cilia or no cilia at all, and no soundevoked potential is evident in these mutant ﬂies.39 The outer segment genes (osegs) constitute a novel family whose members contain multiple tandem tryptophan-aspartate (WD)- and TPR-like repeats.44 The oseg genes were isolated through a phylogenetic screen that identiﬁed genes present in the genomes of ciliated, but not nonciliated, organisms. WD- and TPR-like repeats are found in proteins that form large macromolecular assemblies.44–47 The proteins most closely related to OSEGs are the α’- and β’-coatomers and the clathrin heavy chains, all of which are intimately involved in intracellular trafﬁcking.48–50 Taken together, these observations suggest that OSEGs function as ciliary transport proteins by forming transport complexes. OSEGs are localized at the bases and distal ends of cilia;44 oseg1 and oseg2 mutant ﬂies show abnormal mechanosensory and chemosensory responses and display a severe reduction in or a complete loss of microtubule content at the distal ends of cilia.44 In oseg2 mutant ﬂies, α-tubulin—the main component of cilia and bristles—fails to enter the outer segment of sensory neurons, while this protein is transported normally in oseg1 mutants.44 As mentioned above, kinesin II is the motor for the anterograde IFT and is essential for ciliogenesis as well as for growth and maintenance.51–54 Kinesinassociated protein (KAP) is the nonmotor accessory subunit of kinesin II.55 Mutations in the Drosophila kap gene (DmKap) cause the generation of empty dendritic caps and, in some cases, membranes devoid of axonemes; basal bodies and the ciliary root

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are intact in these mutants.56 Mutations in klp64D, which encodes the kinesin II motor subunit in Drosophila, share certain oseg2 mutant phenotypes, as evidenced by the fact that, in klp64D mutant ﬂies, a green ﬂuorescent protein (GFP)-α-tubulin fusion protein fails to enter the distal ends of cilia and microtubules that accumulate at the base of the cilia.44,56 These ﬁndings suggest that intracellular transporting to the distal portions of cilia requires kinesin-based transport and OSEG proteins. OSEGs and kinesin II are not essential for the synthesis of the proximal ciliary structures.44 The EB1 protein belongs to the family of microtubule plus-end tracking proteins, which are localized at the plus ends of growing microtubules and regulate microtubule dynamics.57–60 In chordotonal organs, the Drosophila EB1 protein (DmEB1) is concentrated in the scolopale region as well as in the area surrounding the inner dendritic segments of sensory neurons. In DmEB1 mutants, the cilia are misaligned and malformed,29 indicating a disruption in the organization of the chordotonal organs. These mutant ﬂies also exhibit uncoordinated movement and a reduced response to sound stimuli.29 These phenotypes indicate that DmEB1 plays an essential role in chordotonal organ structure and function. Mutations in vertebrate myosin VIIA (myoVIIA), an unconventional myosin, lead to deafness, vestibular dysfunction, and retinitis pigmentosa, all of which are characteristics of Usher syndrome Type 1B61 in which the stereocilia of hair bundles are disorganized.61–66 The Drosophila gene that encodes myoVIIA is called crinkled (ck),67 and the loss of Ck function in the Drosophila chordotonal organs leads to complete deafness. This phenotype is likely the result of disruption of the integrity of the scolopidia, including apical detachment, incomplete dendritic capping, and overall scolopidium disorganization.68 The Ck protein is expressed chieﬂy in scolopale rods and scolopale cell-cap cell junctions.68 Gene Products Involved in Auditory Transduction, but Not in External Sensory Transduction Although myriad genes have been identiﬁed that participate in both auditory and external sensory transduction, only four genes—beethoven (btv), touch-insensitivelarval B (tilB), nanchung (nan), and inactive (iav)—are required for the formation or function of chordotonal sensory organs.12,15,16,20 These ﬁndings suggest the existence of properties unique to chordotonal mechanotransduction. Kernan and colleagues isolated tilB using behavioral touch-insensitivity screening.69 Eberl et al.70 isolated btv in a genetic screen that took advantage of the fact that male ﬂies will court one another if they are presented with a courtship song. btv mutant ﬂies display anatomically aberrant ciliary dilation only in the chordotonal cilia.12 The absence of a sound-evoked potential in btv mutant ﬂies indicates that ciliary dilation may have a speciﬁc function in mechanotransduction by the chordotonal cilia.12 tilB mutants have abnormal chordotonal function and produce immotile sperm, but bristle sensory function remains intact.12,69 In spermatids from tilB mutant ﬂies, some of the axonemal proﬁles are split, fragmented, and lack inner and outer dynein arms,12 suggesting that ciliary movement by dynein plays a critical role in chordotonal transduction as well as in ciliary motility. The molecular properties of the Btv and TilB proteins have not yet been deciphered.

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DROSOPHILA TRPV CHANNELS: NANCHUNG AND INACTIVE Ion channels in the TRP superfamily each contain six transmembrane domains along with cytoplasmic N- and C-terminal ends. On the basis of sequence homology, this superfamily was divided into seven subfamilies: TRPC (for classical or canonical), TRPV (for vanilloid-related), TRPM (for melastatin-related), TRPP (for polycystintype), TRPN (for NompA), TRPA (for ANKTM1), and TRPML (for mucolipin). Most TRP family members are nonselective cation channels that function in diverse types of sensory signaling, such as pheromone reception, pain sensation, light perception, gustation, and thermosensation.4,5,71–81 In mammals, the TRPV subfamily has six members.78,81 Patch-clamp analysis has shown that mammalian TRPV channels are gated by a variety of ligands, including capsaicin, certain lipids, various temperatures, and osmotic pressure.4,78,81–84 Genetic analysis revealed that mammalian TRPV channels are required for the production of diverse sensations, including pain, thermosensation, osmosensation, and mechanosensation, as well as for Ca2+ uptake. The worm C. elegans contains ﬁve TRPV channels; the ﬁrst to be identiﬁed was Osm-9 and, later, other family members were discovered, including Ocr-1, 2, 3, and 4 (OSM-9- and capsaicin receptor-related channels 1 to 4).85,86 Osm-9 can form heteromeric complexes with all of the Ocrs. This complex performs speciﬁc functions in selected sensory cells, such as those required for the nose touch sensation, osmolar detection, and olfactory adaptation.86

NANCHUNG

AND INACTIVE

The Drosophila genome contains two genes that encode TRPV channels: nanchung (nan) and inactive (iav).15,16 If one considers protein sequence homology, the Nan protein is most closely related to the C. elegans Ocrs, whereas the Iav protein is most similar to C. elegans Osm-9.4 In heterologous cell types, nan and iav exhibit gating by hypoosmotic stress, suggesting that these channels respond to membrane tension.15,16 However, it is still not known whether second messengers are involved in gating Nan and Iav channels. Both nan and iav display a similar preference for Na+ and K+ ions.4 In situ staining of Drosophila embryos with anti-sense RNA probes shows that both nan and iav are expressed exclusively in the chordotonal organs.15,16 Experiments that assessed the expression of nan promoter-Gal4 gene fusion constructs clearly showed that nan is expressed in neurons. Antisera raised against both the N- and C-terminal regions of the Nan and Iav proteins revealed that nan and iav are localized to the cilia of sensory neurons.15,16 An Iav-GFP fusion construct was localized exclusively in the proximal region of the ciliary segment,16 although the possibility of weak expression in the distal region has not yet been ruled out. Several deletions in the nan locus were generated from the imprecise excision of a P-element located upstream of nan.15 When forced to the bottom of a vial, the resulting mutant ﬂies very quickly climb up the wall and tend to reside near the top surface of the vial, indicating that each ﬂy’s geotaxis—movement against gravity—is

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not impaired. However, the mutant ﬂies show mild uncoordination and restricted movement. In addition, these ﬂies easily fall from the top surface of the vial, which may be indicative of deﬁciencies in proprioception. Similar phenotypes are observed in iav mutant ﬂies, although the effects are more severe relative to the nan mutants.16 The reason for these observed differences is not clear. When assessed for sound-evoked potential, both nan and iav mutant ﬂies show no response, which indicates that these genes are required for mechanotransduction. The ciliary structure and organization appear to be intact, at least in the nan mutants.15 Immunostaining of iav mutant ﬂies with antisera raised against the Nan protein revealed that nan is not detectable anywhere in the sensory neurons.16 The opposite was also true: immunostaining of nan mutant ﬂies with antisera raised against the Iav protein showed that no Iav protein existed anywhere in the neurons. This means that without one channel component, the other failed to escape degradation. This suggests that chordotonal neurons have a quality-control system to ensure that only heteromeric Nan-Iav complexes are localized to the cilia. In addition, these ﬁndings may imply that homomultimeric Nan or Iav complexes do not function in cilia for mechanotransduction. Fleshing out the details of this putative quality-control system constitutes a future challenge.

HOW IS NAN-IAV GATED

IN

VIVO?

Because Nan-Iav heteromeric complexes are required for sound-evoked potential, are located where mechanical force occurs, and are gated by membrane tension generated by hypoosmotic stress, these complexes likely represent transducers that mediate the conversion of mechanical force transmitted to the cilia into neuronal signaling. If Nan-Iav channels are bona ﬁde mechanotransducers, how are the channels gated? The conventional gating model, termed the “trapdoor model,” was proposed to explain the mechanisms of action of hair cell mechanoreceptors and worm touch receptors.8,87 This model postulates that ion channels are anchored between the internal cytoskeleton and the extracellular anchor (Figure 17.3). Movement of one anchor relative to the other would pull the channel open. In the gentle touch response of C. elegans, a member of the DEG/ENaC (degenerin/epithelial sodium channel) transduction channel family is anchored to the intracellular cytoskeleton and extracellular structures via links.8,87–89 The links, cytoskeleton, and several components of the extracellular material were found to constitute the touch transduction machinery in worm touch receptors.6,87,90 In hair cell transduction, a candidate transducer was identiﬁed recently; this protein is expressed in hair cells and localized to the upper region of hair bundles. This channel, called TRPA1, contains 17 ankyrin (ANK) repeats, which have been proposed to form the gating spring.91,92 If chordotonal sensation follows the ‘‘trapdoor model’’ the probable site in which chordotonal transduction would occur is at the ends of the cilia, where the cilia meet the NompA-containing cap structure.33 However, this generalization does not apply to Nan-Iav heteromers, because they are not localized at the cap (Figure 17.1). Instead, they are housed at the proximal ends of the cilia, before the ciliary dilation.16 Therefore, Nan-Iav channels cannot contact NompA. Moreover, extracellular

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FIGURE 17.3 Hypothetical models depicting mechanosensitive channel gating mechanisms. (A) The conventional “trapdoor model.”8,87 Relative displacement of the extracellular anchor and the intracellular cytoskeleton (as shown by the arrow) opens Nan-Iav channels via hypothetical links between the channels and extracellular and intracellular components. (B) The axoneme-dependent bending model. The channel is tightly linked to the microtubulerich axoneme via hypothetical intracellular links. Bending of the axoneme exerts a force (indicated by the arrow), which results in the opening of the channel. Closed circles denote cations. EA, extracellular anchor; EL, extracellular link; IL, intracellular link; CS, cytoskeleton (microtubule axoneme).

anchors presumably are lacking in the proximal region of the cilia. Thus the ‘‘trapdoor model’’ is not applicable to Nan-Iav heteromers. But then how is Nan-Iav gated? I propose an alternative, axoneme-dependent bending model (Figure 17.3). In this scenario, the channel is ﬁrmly attached to the axoneme via links. Rotational movement of the joint in the chordotonal organ would result in the bending of the axoneme of the cilia, which can cause a lateral displacement (via the links) of the channel complexes to the ‘‘open’’ conﬁguration (illustrated in Figure 17.3). The challenge is to determine whether Drosophila chordotonal cilia are bent when mechanical stretch is applied at the ends of the cilia. Indeed, this type of bending is known to occur in the femoral chordotonal organ (FCO) of the grasshopper leg.19 The grasshopper FCO is attached proximally to the dorsal cuticle of the femur and

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distally via ligament cells to the femoro-tibial joint.28 The cilia of the FCO lie slightly off axis from the midline of the ligament, such that pulling the ligament imparts both lateral and longitudinal components of displacement.19 The lateral component is estimated to be an angular deviation of 1 to 10 degrees.19 If the Nan-Iav heteromer is held in a high-energy state, this slight bending might be enough to cause the transduction channel to open. Moreover, greater bending occurs at the proximal part of cilia when the grasshopper tibia is moved.19

CONCLUDING REMARKS Channel properties, subcellular location, and genetic requirements all suggest that the heteromeric Nan-Iav complex functions as a mechanotransducer in the chordotonal organ. On the basis of data reviewed herein, as well as the proximal location of the Nan-Iav heteromer, I propose the axoneme-dependent bending model, in which tight linkage of the Nan-Iav channel to the axoneme is of crucial importance in channel opening when the cilia are bent by mechanical stimulation. The ﬁnding that the Nan-Iav complex is gated in a hypotonic solution would lend support to this model. However, the question of whether or not Nan-Iav is opened directly by membrane expansion needs more scrutiny. Also, additional research is needed to determine whether bending occurs upon stimulation and whether bending is the primary cause of mechanotransduction in the antennal chordotonal organs of the ﬂy.

ACKNOWLEDGMENTS This work was supported in part by a National Research Laboratory (NRL) grant 2005-01335 (Korea Research Foundation) to C.K.

REFERENCES 1. Todi, S.V., Sharma, Y., and Eberl, D.F., Anatomical and molecular design of the Drosophila antenna as a ﬂagellar auditory organ, Microsc. Res. Tech. 63 (6), 388–99, 2004. 2. Boekhoff-Falk, G., Hearing in Drosophila: development of Johnston’s organ and emerging parallels to vertebrate ear development, Dev. Dyn. 232 (3), 550–58, 2005. 3. Corey, D.P., New TRP channels in hearing and mechanosensation, Neuron 39, 585–88, 2003. 4. Liedtke, W. and Kim, C., Functionality of the TRPV subfamily of TRP ion channels: add mechano-TRP and osmo-TRP to the lexicon! Cell Mol. Life Sci. 62 (24), 2985–3001, 2005. 5. Lin, S.Y. and Corey, D.P., TRP channels in mechanosensation, Curr. Opin. Neurobiol. 15 (3), 350–57, 2005. 6. Gillespie, P.G., Dumont, R.A., and Kachar, B., Have we found the tip link, transduction channel, and gating spring of the hair cell? Curr. Opin. Neurobiol. 15 (4), 389–96, 2005. 7. Sukharev, S. and Corey, D.P., Mechanosensitive channels: multiplicity of families and gating paradigms, Sci. STKE 2004 (219), re4, 2004.

238

TRP Ion Channel Function in Sensory Transduction

8. Kung, C., A possible unifying principle for mechanosensation, Nature 436 (7051), 647–54, 2005. 9. Jarman, A.P., Studies of mechanosensation using the ﬂy, Hum. Mol. Genet. 11 (10), 1215–18, 2002. 10. Bermingham, N.A., Hassan, B.A., Price, S.D., Vollrath, M.A., Ben-Arie, N., Eatock, R.A., Bellen, H.J., Lysakowski, A., and Zoghbi, H.Y., Math1: an essential gene for the generation of inner ear hair cells, Science 284 (5421), 1837–41, 1999. 11. Jarman, A.P., Grau, Y., Jan, L.Y., and Jan, Y.N., atonal is a proneural gene that directs chordotonal organ formation in the Drosophila peripheral nervous system, Cell 73 (7), 1307–21, 1993. 12. Eberl, D.F., Hardy, R.W., and Kernan, M.J., Genetically similar transduction mechanisms for touch and hearing in Drosophila, J. Neurosci. 20 (16), 5981–88, 2000. 13. Wang, V.Y., Hassan, B.A., Bellen, H.J., and Zoghbi, H.Y., Drosophila atonal fully rescues the phenotype of Math1 null mice: new functions evolve in new cellular contexts, Curr. Biol. 12 (18), 1611–16, 2002. 14. Ben-Arie, N., Hassan, B.A., Bermingham, N.A., Malicki, D.M., Armstrong, D., Matzuk, M., Bellen, H.J., and Zoghbi, H.Y., Functional conservation of atonal and Math1 in the CNS and PNS, Development 127 (5), 1039–48, 2000. 15. Kim, J., Chung, Y.D., Park, D.Y., Choi, S., Shin, D.W., Soh, H., Lee, H.W., Son, W., Yim, J., Park, C.S., Kernan, M.J., and Kim, C., A TRPV family ion channel required for hearing in Drosophila, Nature 424 (6944), 81–84, 2003. 16. Gong, Z., Son, W., Chung, Y.D., Kim, J., Shin, D.W., McClung, C.A., Lee, Y., Lee, H.W., Chang, D.J., Kaang, B.K., Cho, H., Oh, U., Hirsh, J., Kernan, M.J., and Kim, C., Two interdependent TRPV channel subunits, inactive and Nanchung, mediate hearing in Drosophila, J. Neurosci. 24 (41), 9059–66, 2004. 17. Corey, D.P., García-Añoveros, J., Holt, J.R., Kwan, K.Y., Lin, S.Y., Vollrath, M.A., Amalﬁtano, A., Cheung, E.L., Derﬂer, B.H., Duggan, A., Geleoc, G.S., Gray, P.A., Hoffman, M.P., Rehm, H.L., Tamasauskas, D., and Zhang, D.S., TRPA1 is a candidate for the mechanosensitive transduction channel of vertebrate hair cells, Nature 432 (7018), 723–30, 2004. 18. Burns, M.D., Structure and physiology of the locust femoral chordotonal organ, J. Insect Physiol. 20 (7), 1319–39, 1974. 19. Moran, D.T., Varela, F.J., and Rowley, J.C., 3rd, Evidence for active role of cilia in sensory transduction, Proc. Natl. Acad. Sci. USA 74 (2), 793–97, 1977. 20. Eberl, D.F., Feeling the vibes: chordotonal mechanisms in insect hearing, Curr. Opin. Neurobiol. 9 (4), 389–93, 1999. 21. Caldwell, J.C. and Eberl, D.F., Towards a molecular understanding of Drosophila hearing, J. Neurobiol. 53 (2), 172–89, 2002. 22. Robert, D. and Gopfert, M.C., Novel schemes for hearing and orientation in insects, Curr. Opin. Neurobiol. 12 (6), 715–20, 2002. 23. Bennet-Clark, H.C., Acoustics of insect song, Nature 234, 255–59, 1971. 24. Hall, J.C., The mating of a ﬂy, Science 264 (5166), 1702–14, 1994. 25. Greenspan, R.J. and Ferveur, J.F., Courtship in Drosophila, Annu. Rev. Genet. 34, 205–32, 2000. 26. Tauber, E. and Eberl, D.F., Acoustic communication in Drosophila, Behav. Processes 64 (2), 197–210, 2003. 27. Moulins, P.J., Structure and function of proprioceptors in the invertebrates. Chapman and Hall, London, 1976.

TRPV Family Ion Channels

239

28. Moran, D.T., Rowley, J.C., 3rd, and Varela, F.G., Ultrastructure of the grasshopper proximal femoral chordotonal organ, Cell Tissue Res. 161 (4), 445–57, 1975. 29. Elliott, S.L., Cullen, C.F., Wrobel, N., Kernan, M.J., and Ohkura, H., EB1 is essential during Drosophila development and plays a crucial role in the integrity of chordotonal mechanosensory organs, Mol. Biol. Cell 16 (2), 891–901, 2005. 30. Moran, D.T. and Rowley, J.C., 3rd, The ﬁne structure of the cockroach subgenual organ, Tissue Cell 7 (1), 91–105, 1975. 31. Baker, J.D., Adhikarakunnathu, S., and Kernan, M.J., Mechanosensory-defective, male-sterile unc mutants identify a novel basal body protein required for ciliogenesis in Drosophila, Development 131 (14), 3411–22, 2004. 32. Martinez-Campos, M., Basto, R., Baker, J., Kernan, M., and Raff, J.W., The Drosophila pericentrin-like protein is essential for cilia/ﬂagella function, but appears to be dispensable for mitosis, J. Cell Biol. 165 (5), 673–83, 2004. 33. Chung, Y.D., Zhu, J., Han, Y., and Kernan, M.J., nompA encodes a PNS-speciﬁc, ZP domain protein required to connect mechanosensory dendrites to sensory structures, Neuron 29 (2), 415–28, 2001. 34. Bork, P. and Sander, C., A large domain common to sperm receptors (Zp2 and Zp3) and TGF-beta type III receptor, FEBS Lett. 300 (3), 237–40, 1992. 35. Li, X.J. and Snyder, S.H., Molecular cloning of Ebnerin, a von Ebner’s gland protein associated with taste buds, J. Biol. Chem. 270 (30), 17674–79, 1995. 36. Tordai, H., Banyai, L., and Patthy, L., The PAN module: the N-terminal domains of plasminogen and hepatocyte growth factor are homologous with the apple domains of the prekallikrein family and with a novel domain found in numerous nematode proteins, FEBS Lett. 461 (1–2), 63–67, 1999. 37. Legan, P.K., Rau, A., Keen, J.N., and Richardson, G.P., The mouse tectorins. Modular matrix proteins of the inner ear homologous to components of the sperm-egg adhesion system, J. Biol. Chem. 272 (13), 8791–801, 1997. 38. Mustapha, M., Weil, D., Chardenoux, S., Elias, S., El-Zir, E., Beckmann, J.S., Loiselet, J., and Petit, C., An alpha-tectorin gene defect causes a newly identiﬁed autosomal recessive form of sensorineural pre-lingual non-syndromic deafness, DFNB21, Hum. Mol. Genet. 8 (3), 409–12, 1999. 39. Han, Y.G., Kwok, B.H., and Kernan, M.J., Intraﬂagellar transport is required in Drosophila to differentiate sensory cilia but not sperm, Curr. Biol. 13 (19), 1679–86, 2003. 40. Kozminski, K.G., Beech, P.L., and Rosenbaum, J.L., The Chlamydomonas kinesinlike protein FLA10 is involved in motility associated with the ﬂagellar membrane, J. Cell Biol. 131 (6 Pt. 1), 1517–27, 1995. 41. Huang, B., Rifkin, M.R., and Luck, D.J., Temperature-sensitive mutations affecting ﬂagellar assembly and function in Chlamydomonas reinhardtii, J. Cell Biol. 72 (1), 67–85, 1977. 42. Marshall, W.F. and Rosenbaum, J.L., Intraﬂagellar transport balances continuous turnover of outer doublet microtubules: implications for ﬂagellar length control, J. Cell Biol. 155 (3), 405–14, 2001. 43. Rosenbaum, J.L. and Witman, G.B., Intraﬂagellar transport, Nat. Rev. Mol. Cell Biol. 3 (11), 813–25, 2002. 44. Avidor-Reiss, T., Maer, A.M., Koundakjian, E., Polyanovsky, A., Keil, T., Subramaniam, S., and Zuker, C.S., Decoding cilia function: deﬁning specialized genes required for compartmentalized cilia biogenesis, Cell 117 (4), 527–39, 2004.

240

TRP Ion Channel Function in Sensory Transduction

45. Das, A.K., Cohen, P.W., and Barford, D., The structure of the tetratricopeptide repeats of protein phosphatase 5: implications for TPR-mediated protein–protein interactions, Embo J. 17 (5), 1192–99, 1998. 46. Cronshaw, J.M. and Matunis, M.J., The nuclear pore complex protein ALADIN is mislocalized in triple A syndrome, Proc. Natl. Acad. Sci. USA 100 (10), 5823–27, 2003. 47. Gotzmann, J., Gerner, C., Meissner, M., Holzmann, K., Grimm, R., Mikulits, W., and Sauermann, G., hNMP 200: a novel human common nuclear matrix protein combining structural and regulatory functions, Exp. Cell Res. 261 (1), 166–79, 2000. 48. Kirchhausen, T., Three ways to make a vesicle, Nat. Rev. Mol. Cell Biol. 1 (3), 187–98, 2000. 49. Perrais, D. and Merriﬁeld, C.J., Dynamics of endocytic vesicle creation, Dev. Cell 9 (5), 581–92, 2005. 50. McPherson, P.S. and Ritter, B., Peptide motifs: building the clathrin machinery, Mol. Neurobiol. 32 (1), 73–87, 2005. 51. Marszalek, J.R. and Goldstein, L.S., Understanding the functions of kinesin-II, Biochim. Biophys. Acta. 1496 (1), 142–50, 2000. 52. Ou, G., Blacque, O.E., Snow, J.J., Leroux, M.R., and Scholey, J.M., Functional coordination of intraﬂagellar transport motors, Nature 436 (7050), 583–87, 2005. 53. Snow, J.J., Ou, G., Gunnarson, A.L., Walker, M.R., Zhou, H.M., Brust-Mascher, I., and Scholey, J.M., Two anterograde intraﬂagellar transport motors cooperate to build sensory cilia on C. elegans neurons, Nat. Cell Biol. 6 (11), 1109–13, 2004. 54. Scholey, J.M., Intraﬂagellar transport, Annu. Rev. Cell Dev. Biol. 19, 423–43, 2003. 55. Sarpal, R. and Ray, K., Dynamic expression pattern of kinesin accessory protein in Drosophila, J. Biosci. 27 (5), 479–87, 2002. 56. Sarpal, R., Todi, S.V., Sivan-Loukianova, E., Shirolikar, S., Subramanian, N., Raff, E.C., Erickson, J.W., Ray, K., and Eberl, D.F., Drosophila KAP interacts with the kinesin II motor subunit KLP64D to assemble chordotonal sensory cilia, but not sperm tails, Curr. Biol. 13 (19), 1687–96, 2003. 57. Tirnauer, J.S. and Bierer, B.E., EB1 proteins regulate microtubule dynamics, cell polarity, and chromosome stability, J. Cell Biol. 149 (4), 761–66, 2000. 58. Vaughan, K.T., Microtubule plus ends, motors, and trafﬁc of Golgi membranes, Biochim. Biophys. Acta. 1744 (3), 316–24, 2005. 59. Grevengoed, E.E. and Peifer, M., Cytoskeletal connections: building strong cells in new ways, Curr. Biol. 13 (14), R568–70, 2003. 60. Schuyler, S.C. and Pellman, D., Microtubule ‘‘plus-end-tracking proteins’’: The end is just the beginning, Cell 105 (4), 421–24, 2001. 61. El-Amraoui, A. and Petit, C., Usher I syndrome: unravelling the mechanisms that underlie the cohesion of the growing hair bundle in inner ear sensory cells, J. Cell Sci. 118 (Pt. 20), 4593–603, 2005. 62. Rzadzinska, A.K., Schneider, M.E., Davies, C., Riordan, G.P., and Kachar, B., An actin molecular treadmill and myosins maintain stereocilia functional architecture and self-renewal, J. Cell Biol. 164 (6), 887–97, 2004. 63. Boeda, B., El-Amraoui, A., Bahloul, A., Goodyear, R., Daviet, L., Blanchard, S., Perfettini, I., Fath, K.R., Shorte, S., Reiners, J., Houdusse, A., Legrain, P., Wolfrum, U., Richardson, G., and Petit, C., Myosin VIIa, harmonin, and cadherin 23, three Usher I gene products that cooperate to shape the sensory hair cell bundle, Embo J. 21 (24), 6689–99, 2002. 64. Holme, R.H. and Steel, K.P., Stereocilia defects in waltzer (Cdh23), shaker1 (Myo7a) and double waltzer/shaker1 mutant mice, Hear. Res. 169 (1–2), 13–23, 2002.

TRPV Family Ion Channels

241

65. Keats, B.J. and Corey, D.P., The usher syndromes, Am. J. Med. Genet. 89 (3), 158–66, 1999. 66. Weil, D., Levy, G., Sahly, I., Levi-Acobas, F., Blanchard, S., El-Amraoui, A., Crozet, F., Philippe, H., Abitbol, M., and Petit, C., Human myosin VIIA responsible for the Usher 1B syndrome: a predicted membrane-associated motor protein expressed in developing sensory epithelia, Proc. Natl. Acad. Sci. USA 93 (8), 3232–37, 1996. 67. Kiehart, D.P., Franke, J.D., Chee, M.K., Montague, R.A., Chen, T.L., Roote, J., and Ashburner, M., Drosophila crinkled, mutations of which disrupt morphogenesis and cause lethality, encodes ﬂy myosin VIIA, Genetics 168 (3), 1337–52, 2004. 68. Todi, S.V., Franke, J.D., Kiehart, D.P., and Eberl, D.F., Myosin VIIA defects, which underlie the Usher 1B syndrome in humans, lead to deafness in Drosophila, Curr. Biol. 15 (9), 862–68, 2005. 69. Kernan, M., Cowan, D., and Zuker, C., Genetic dissection of mechanosensory transduction: mechanoreception-defective mutations of Drosophila, Neuron 12 (6), 1195–206, 1994. 70. Eberl, D.F., Duyk, G.M., and Perrimon, N., A genetic screen for mutations that disrupt an auditory response in Drosophila melanogaster, Proc. Natl. Acad. Sci. USA 94 (26), 14837–42, 1997. 71. Ramsey, I.S., Delling, M., and Clapham, D.E., An introduction to TRP channels, Annu. Rev. Physiol., 2005. 72. Pedersen, S.F., Owsianik, G., and Nilius, B., TRP channels: an overview, Cell Calcium 38 (3–4), 233–52, 2005. 73. Lee, H. and Caterina, M.J., TRPV channels as thermosensory receptors in epithelial cells, Pﬂügers Arch. 451 (1), 160–67, 2005. 74. Delmas, P., Polycystins: polymodal receptor/ion-channel cellular sensors, Pﬂügers Arch. 451 (1), 264–76, 2005. 75. Tominaga, M. and Caterina, M.J., Thermosensation and pain, J. Neurobiol. 61 (1), 3–12, 2004. 76. Vriens, J., Owsianik, G., Voets, T., Droogmans, G., and Nilius, B., Invertebrate TRP proteins as functional models for mammalian channels, Pﬂügers Arch. 449 (3), 213–26, 2004. 77. Moran, M.M., Xu, H., and Clapham, D.E., TRP ion channels in the nervous system, Curr. Opin. Neurobiol. 14 (3), 362–69, 2004. 78. Mutai, H. and Heller, S., Vertebrate and invertebrate TRPV-like mechanoreceptors, Cell Calcium 33 (5–6), 471–78, 2003. 79. Patapoutian, A., TRP channels and thermosensation, Chem. Senses 30 Suppl. 1, i193–i194, 2005. 80. Voets, T. and Nilius, B., TRPs make sense, J. Membr. Biol. 192 (1), 1–8, 2003. 81. Montell, C., The TRP superfamily of cation channels, Sci. STKE 2005 (272), re3, 2005. 82. Benham, C.D., Gunthorpe, M.J., and Davis, J.B., TRPV channels as temperature sensors, Cell Calcium 33 (5–6), 479–87, 2003. 83. Nilius, B. and Voets, T., Diversity of TRP channel activation, Novartis Found. Symp. 258, 140–49; discussion 149–59, 263–66, 2004. 84. Nilius, B., Vriens, J., Prenen, J., Droogmans, G., and Voets, T., TRPV4 calcium entry channel: a paradigm for gating diversity, Am. J. Physiol. Cell Physiol. 286 (2), C195–205, 2004. 85. Colbert, H.A., Smith, T.L., and Bargmann, C.I., OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans, J. Neurosci. 17 (21), 8259–69, 1997.

242

TRP Ion Channel Function in Sensory Transduction

86. Tobin, D., Madsen, D.M., Kahn-Kirby, A., Peckol, E., Moulder, G., Barstead, R., Maricq, A.V., and Bargmann, C.I., Combinatorial expression of TRPV channel proteins deﬁnes their sensory functions and subcellular localization in C. elegans neurons, Neuron 35, 307–18, 2002. 87. Gillespie, P.G. and Walker, R.G., Molecular basis of mechanosensory transduction, Nature 413 (6852), 194–202, 2001. 88. Goodman, M.B. and Schwarz, E.M., Transducing touch in Caenorhabditis elegans, Annu. Rev. Physiol. 65, 429–52, 2003. 89. Hamill, O.P. and Martinac, B., Molecular basis of mechanotransduction in living cells, Physiol. Rev. 81 (2), 685–740, 2001. 90. LeMasurier, M. and Gillespie, P.G., Hair-cell mechanotransduction and cochlear ampliﬁcation, Neuron 48 (3), 403–15, 2005. 91. Sotomayor, M., Corey, D.P., and Schulten, K., In search of the hair-cell gating spring elastic properties of ankyrin and cadherin repeats, Structure 13 (4), 669–82, 2005. 92. Howard, J. and Bechstedt, S., Hypothesis: a helix of ankyrin repeats of the NOMPCTRP ion channel is the gating spring of mechanoreceptors, Curr. Biol. 14 (6), R224–26, 2004.

18

The TRPV Channel in C. elegans Serotonergic Neurons Ji Ying Sze University of California, Irvine

CONTENTS Abstract ..................................................................................................................243 Introduction............................................................................................................244 Conservation of the Serotonergic System in C. elegans ......................................244 The Function and Mechanism of OCR-2 and OSM-9 in Serotonergic Neurons ........................................................................................246 Speciﬁcity of OCR-2 and OSM-9 Signaling Pathways in Different Sensory Neurons................................................................................247 Sensory Receptors .....................................................................................247 Components in Signaling Cascades ..........................................................249 Intrinsic Modality Determinant .................................................................249 TRP Channels in Mammalian Serotonergic Systems? .........................................251 References..............................................................................................................252

ABSTRACT In the pair of the nematode Caenorhabiditis elegans serotonergic chemosensory neurons ADF, the TRPV channel protein OCR-2 interacts with another TRPV protein, OSM-9, to control the production of the neurotransmitter serotonin. The activity and speciﬁcity of OCR-2 in the serotonergic neurons is governed by structural determinants within the channel protein in concert with deﬁned cellular components. The dynamic gating mechanisms, multiple sensory modalities, and functional conservation in diverse organisms make TRPV channels ideal candidates for the long-awaited molecular sensors that underscore the ancient role of the serotonergic system in coupling sensory cues and internal milieu to behavior and physiology.

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INTRODUCTION The serotonergic system is an ancient sensor of diverse stimuli. Serotonin (5-hydroxytryptamine; 5-HT) is a monoamine and functions as a neuromodulator participating in the elaboration of adaptive responses to external stimuli and physiological challenges. The ﬁrst demonstration of serotonin in regulating sensory response was from studies in Aplysia, where increased serotonin promotes sensitization of the gill-withdrawal reﬂex in response to repetitive tactile stimulus to the mantle or the edge of the siphon (Brunelli et al., 1976; Bailey et al., 1992). Although a serotonin signal is produced from the interneurons rather than from the sensory neurons in the gill-withdrawal reﬂex circuitry (Kandel and Schwartz, 1982), this showed that the activity of a speciﬁc set of serotonergic neurons is modulated by a speciﬁc external cue to inﬂuence behavior. Anatomic analysis of the serotonergic system in mammalian CNS revealed their close relationship to blood vessels, leading to the proposal that the raphe serotonergic neurons may directly function as chemoreceptors and mechanoreceptors to transduce diverse neuronal and nonneuronal signals into discrete behavior and physiological processes (Scheibel et al., 1975; Azmitia, 1999). Consistent with this notion, application of distinct chemical and physical stimuli to whole organisms or brain slices induces up- and downregulation of serotonin synthesis in discrete serotonergic neurons in the CNS (Boadle-Biber, 1993; Leibowitz and Alexander, 1998; Azmitia, 1999; Chaouloff et al., 1999; Adell et al., 2002). If serotonergic neurons can discriminate ligands or cues that reﬂect external stimuli and internal milieu, molecular biology predicts that different serotonergic neurons express different receptors. Two such cell-speciﬁc regulators have been identiﬁed in C. elegans: the TRPV (vanilloid subfamily of transient receptor potential) channel proteins OCR-2 and OSM-9. This chapter highlights the conservation of the serotonergic system in C. elegans, describes the role of OCR-2 and OSM-9 in the serotonergic neurons, explains how these two TRPV channel proteins can be activated by multiple sensory stimuli but maintain speciﬁcity to each modality, and, ﬁnally, discusses the possibility of serotonin as a conserved readout of TRP channel signaling.

CONSERVATION OF THE SEROTONERGIC SYSTEM IN C. ELEGANS As in all animals, serotonergic neurons represent a small population in C. elegans. A mature hermaphroditic worm has exactly 302 neurons (White et al., 1986). The nine neurons that can be stained by an antibody raised against serotonin are composed of ﬁve neural classes, with four classes consisting of two bilaterally symmetric neurons (Horvitz et al., 1982; Sze et al., 2000). C. elegans does not have a deﬁned CNS; nevertheless, the serotonergic neurons are connected to the central neuronal integration center called the nerve ring (White et al., 1986) and inﬂuence sensory behavior, both innate and learned, and diverse physiological processes (for examples, see Weinshenker et al., 1995; Colbert and Bargmann, 1997; Nurrish et al., 1999; Sze et al., 2000; Sawin et al., 2000; Zhang et al., 2005). ADF are the only serotonergic sensory neurons in the animal. The sensory cilia of ADF are exposed to the external

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environment (White et al., 1986) and sense salts (Bargmann and Horvitz, 1991; Hukema et al., 2006). A recent study showed that serotonin produced from ADF mediates olfactory learning (Zhang et al., 2005). NSM are secretory neurons with their cell bodies located in the pharynx and are likely sensing signals released from food (Sawin et al., 2000). HSN are motor neurons that couple feeding status and the rhythm of vulva muscle contractions to release fertilized eggs (Desai et al., 1988). RIH and AIM are interneurons, bridging between sensory inputs and command outputs (White et al., 1986). The unique, distinguishable function of the serotonergic neurons in C. elegans provides a genetic tractable model to investigate the genes that regulate serotonergic phenotypes in identiﬁed neural types. Classic mutant screens in conjunction with gene knockout technology have led to the isolation of mutant animals with deﬁcits in serotonin biosynthesis, serotonin release, and serotonin receptors (Duerr et al., 1999; Sze et al., 2000; Ranganathan et al., 2000; Hare and Loer, 2004; Dempsey et al., 2005). Genes used to produce serotonergic phenotypes are highly conserved between worms and humans. Three of those—tph-1, cat-1, and mod-5—are relevant here. The tph-1 gene encodes the tryptophan hydroxylase that catalyzes the rate-limiting ﬁrst step of serotonin biosynthesis, and the tph-1 deletion mutant has no detectable serotonin (Sze et al., 2000). Like in mammals, signaling by serotonin in C. elegans involves two distinct mechanisms of serotonin transporter. cat-1 encodes the vesicular monoamine transporter (VMAT) on the secretory vesicles that pump newly synthesized serotonin into vesicles for regulated exocytosis (Duerr et al., 1999), and mod-5 encodes the membrane serotonin transporter (SERT) for reuptake of extracellular serotonin (Ranganathan et al., 2001). SERT is the target of two major classes of antidepressants: selective serotonin reuptake inhibitors (SSRIs) and tricyclic drugs. It is generally assumed that these drugs exert therapeutic effects by blocking SERT from reuptaking serotonin, thereby increasing the availability of serotonin at the synapse (Blier and de Montigny, 1998). A recent study in our laboratory found that worms either bearing a deletion in the SERT gene mod-5, or wild-type worms treated with the SSRI ﬂuoxetine, accumulate serotonin only in the ADF, NSM, and HSN neurons (C. Dempsey and J. Sze, unpublished). This indicates that two classes of the serotonergic neurons in the head (AIM and RIH) can only absorb extracellular serotonin via MOD-5/SERT and that ﬂuoxetine blocks MOD-5 activity. Such serotonin-absorbing neurons have also been observed in developing rat embryos throughout sensory pathways from the CNS to sensory neurons (Hansson et al., 1998; Lebrand et al., 1998), and in the hypothalamic dorsomedial nucleus in postnatal life (Hoffman et al., 1998). In worms as well as in mammals, these neurons do not express the complete set of serotonin synthesis enzymes, but they do express VMAT. One possible function for these neurons could be serving as “relay stations” that pass serotonin from originating neuron sources to speciﬁc serotonin receptor subtypes on distant targets. The potential effects of the blockage of SERT by SSRIs and tricyclic drugs in the pathways mediated by these neurons are fascinating but beyond the scope of this chapter. What is relevant is the conservation of the mechanisms of serotonin neurotransmission and of the principle action of the SSRIs on SERT in the worm, which gives a more concrete reason to use the C. elegans serotonergic

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system as a model for understanding the fundamental mechanisms that regulate serotonin signaling.

THE FUNCTION AND MECHANISM OF OCR-2 AND OSM-9 IN SEROTONERGIC NEURONS A role of TRPV channels in serotonin signaling was identiﬁed from an unbiased genetic screen for serotonin synthesis mutants (Zhang et al., 2004). Because tryptophan hydroxylase is the key enzyme for serotonin synthesis, and the expression of GFP tagged the tph-1 gene (tph-1::gfp), which can be unambiguously identiﬁed and, to some extent, quantiﬁed in speciﬁc serotonergic neurons of living worms, a goal of this screen was to ask whether tph-1 expression in different classes of neurons is regulated by different genes. It turned out most mutants retrieved from the screen were missing tph-1::gfp expression speciﬁcally in one class of the serotonergic neurons. The ﬁrst two genes identiﬁed that regulate tph-1 expression in the ADF chemosensory neurons are the TRPV genes osm-9 and ocr-2. Null mutations in either osm-9 or ocr-2 dramatically downregulate tph-1::gfp expression in the ADF neurons, but tph-1::gfp expression in the NSM and HSN neurons is unaffected in the mutants. And there is no detectable difference in the intensity of tph-1::gfp expression between double mutants of ocr-2 and osm-9 with each of the single mutants. Several lines of evidence indicate that both OCR-2 and OSM-9 act in the ADF neurons to regulate serotonin production. OCR-2 and OSM-9 are coexpressed in ADF (Tobin et al., 2002; Zhang et al., 2004). By expressing wild-type ocr-2 or osm-9 coding sequences under cell-speciﬁc promoters, it was demonstrated that expression of OCR-2 and OSM-9 in ADF is necessary and sufﬁcient to restore tph-1 expression in respective mutants. These ﬁndings are consistent with earlier evidence that OCR-2 and OSM-9 require each other for routing to the sensory cilia of the neurons (Tobin et al., 2002), suggesting an OCR-2 and OSM-9 protein complex at the sensory cilia of ADF that regulate tph-1 expression. For the rest of this chapter, the term OCR-2/OSM-9 will be used when they are considered as one complex. Does OCR-2/OSM-9 represent a molecular sensor that couples sensory cues to serotonin signaling? To address this question, it is important to consider how speciﬁc tph-1 downregulation is in the mutants. Biophysical characterization of OCR-2 and OSM-9 in a heterologous expression system has not been successful, but TRP channels are generally considered as calcium-permeable cation channels (Montell, 2005; Pedersen et al., 2005; Ramsey et al., 2006). Disruption of cellular calcium homeostasis can affect a battery of cellular events, even causing cell death (Berridge et al., 2003). However, ocr-2 and osm-9 mutations do not appear to cause cell-fate transformation. In addition to ADF, OSM-9 and OCR-2 are coexpressed in ﬁve pairs of nonserotonergic chemosensory neurons: AWA, ADL, ASH in the head, and PHA and PHB in the tail (Tobin et al., 2002). Mutations in osm-9 and ocr-2 do not cause these neurons to degenerate or to exhibit a morphological change detectable at the level of ﬂuorescence microscopy (Colbert et al., 1997; Zhang et al., 2004). Null mutations in osm-9 and ocr-2 also do not reduce the expression of every gene in ADF.

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In fact, all the ADF neuronal markers that have been examined showed no distinguishable difference between mutants and wild-type animals, including the expression of the cat-1/VMAT gene and the cat-4 gene that encodes the tryptophan hydroxylase cofactor GTP-cyclohydrolase I (Zhang et al., 2004). These ﬁndings suggest that the signaling from OCR-2/OSM-9 selectively, if not speciﬁcally, regulates tph-1 expression. An attractive model is that the TRPV channel is regulated by environmental signals to in turn regulate serotonin signaling by transcriptional regulation of the key serotonin synthetic enzyme tryptophan hydroxylase. Characterization of OCR-2/OSM-9 function in other chemosensory neurons provides some consensus why tph-1 is a selected target of the channel in the ADF neurons. ocr-2 and osm-9 mutants also are defective in olfactory sensation to the odorant diacetyl. Diacetyl is sensed by the G-protein-coupled receptor ODR-10, which is expressed speciﬁcally in the AWA neurons (Sengupta et al., 1996). odr-10 expression but not several other AWA markers is reduced in ocr-2 and osm-9 mutants (Colbert et al., 1997; Tobin et al., 2002). It seems that OCR-2/OSM-9 signaling regulates the expression of genes unique for sensory detection and sensory output of the particular cell. Such activity-dependent transcriptional regulation of sensory components may endow the TRPV channel to regulate the sensitivity to sensory stimuli and underscore the mechanisms of serotonin signaling in modulating behavior based on the sensory experience.

SPECIFICITY OF OCR-2 AND OSM-9 SIGNALING PATHWAYS IN DIFFERENT SENSORY NEURONS OCR-2 and OSM-9 are colocalized in the sensory cilia and plasma membrane of four pairs of chemosensory neurons: ADF, AWA, ASH, and ADL. The sensory cues and activation mechanisms of OCR-2/OSM-9 in ADF are not yet determined. Nevertheless, genetic analyses of OCR-2 and OSM-9 in individual sensory neuron functions and their functional interactions with other components in the sensory signaling pathways have produced some basic ideas for understanding and further deducing how the same channel entity mediates multiple sensory modalities but remains faithful to each sensory function in vivo (Figure 18.1).

SENSORY RECEPTORS One mechanism that dictates OCR-2/OSM-9 modality appears to be G-protein signaling. As stated above, the odorant diacetyl is sensed by the G-protein-coupled receptor ODR-10, which is speciﬁcally expressed in the AWA neurons. Thus, it is likely that the activity of OCR-2/OSM-9 in AWA is regulated by ODR-10 and receptor-coupled signaling cascades. There is compelling genetic evidence to suggest that OCR-2/OSM-9 directly senses environmental osmotic strength and tactile stimuli at the nose through the ASH neurons (Liedtke et al., 2003). However, the function of OCR-2/OSM-9 in osmotic and mechanical sensation also depends on the G protein ODR-3 (Roayaie et al., 1998). This suggests that the OCR-2/OSM-9 function in osmotic and mechanical sensation is speciﬁed by a compound activation of mechanical gating and G-protein signaling. A mammalian TRPV4 can substitute OSM-9 to

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FIGURE 18.1 (Color ﬁgure follows p. 234.) Polymodality and speciﬁcity of OCR-2/OSM-9 in vivo. (A) OCR-2/OSM-9 in different sensory functions are regulated by distinct mechanisms. OCR-2 and OSM-9 likely form a heteromeric channel and are located in the sensory cilia and plasma membrane of four classes of chemosensory neurons in the sensory organ amphids. The membrane region is shown in gray. The transmembrane domains of OCR-2 and OSM-9 are shown as oval columns, the rectangular bar represents the ankyrin motifs, and the ball structure represents the N-terminal region preceding the ankyrin motifs. OCR-2/OSM-9 function in diacetyl sensation depends on the G-protein-coupled receptor ODR-10, the G protein ODR-3, and the signals from polyunsaturated fatty acids, but is not affected by the OCR2(G36E) mutation in the ball. OCR-2/OSM-9 sensation to external osmolarity and gentle touch at the nose requires the G protein ODR-3, the signals from polyunsaturated fatty acids, and the determinants located in the ball region of OCR-2. Upregulation of tph-1 expression in the ADF neurons is governed by the determinants located in the ball of OCR-2. Abbreviations: G, heterotrimeric G protein; GPCR, G-protein-coupled receptor. (B) Photomicrographs of the serotonergic chemosensory neurons ADF expressing OCR-2. Both the wild-type OCR-2 and the mutant OCR-2(G36E) proteins are tagged to the FLAG epitope, and the transgenic worms were stained with anti-FLAG antibody. Notice that both wild-type OCR-2 and OCR-2(G36E) are expressed in the cilia (arrowheads) and the cell bodies.

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direct osmotic and mechanical sensation in C. elegans (Liedtke et al., 2003). The TRPV4 operates via mechanical gating as well as chemical activation in heterologous expression systems (Liedtke et al., 2000; Watanabe et al., 2002; Vriens et al., 2004), suggesting conservation of the function and gating mechanism between worms and mammals.

COMPONENTS

IN

SIGNALING CASCADES

Besides ocr-2 and osm-9, several essential components involved in diacetyl sensation and osmotic sensation have been identiﬁed by genetic approaches. Phenotypic analysis of these mutants revealed components that are required for OCR-2/OSM-9 function in AWA for diacetyl sensation and in ASH for osmotic and mechanical sensation, but they are not required for OCR-2/OSM-9 to upregulate tph-1 expression in ADF. The Gα protein ODR-3 is expressed in AWA, ASH, and ADF (Roayaie et al., 1998). odr-3 mutants are severely defective in response to diacetyl, osmolarity, and gentle touch to the nose (Roayaie et al., 1998), and normal ODR-3 activity is required for the rat TRPV4 to direct osmotic and tactile sensation (Liedtke et al., 2003). However, odr-3 mutations or deletion mutations in any other Gα proteins that are expressed in ADF have no detectable effects on tph-1 expression and serotonin synthesis (Zhang et al., 2004). A recent study found that polyunsaturated fatty acids produced by the fat-3 gene regulate OCR-2/OSM-9 function in diacetyl, osmotic, and mechanical sensation (Kahn-Kirby et al., 2004). Again, tph-1 expression is unaffected in fat-3 deletion mutants (I. Sokolchik and J. Sze, unpublished). These ﬁndings suggest that OCR-2/OSM-9 is assembled to a distinct signaling pathway to regulate serotonin production. That OCR-2/OSM-9 drives distinct signaling cascades is supported by studies of the channel functions in the ASH neurons. OCR-2/OSM-9 mediates at least three sensory modalities through the ASH neurons: high osmolarity, gentle touch at the nose, and noxious olfactory stimuli. Sensation of noxious chemicals induces C. elegans worms to aggregate; the function of OCR-2/OSM-9 in this nociceptive sensation, unlike its activity in osmotic and mechanical sensation, is independent of ODR-3 (de Bono et al., 2002). Furthermore, the rat TRPV4 can restore osmotic and mechanical sensation of osm-9 mutants, but it cannot substitute for OSM-9 to transduce noxious odorant stimuli (Liedtke et al., 2003). These results reinforce the notion derived from studies in Drosophila and mammals that TRP channels are assembled to distinct macromolecular signaling complexes to transduce distinct sensory cues (Tsunoda et al., 1997; Ramsey et al., 2006). The selective sensory functions of the rat TRPV4 in the ASH neurons suggest that the ability for a TRP channel to be integrated into sensory signaling complexes is dictated by structural determinants intrinsic to the channel protein, and these determinants may be selectively conserved and segregated through evolution.

INTRINSIC MODALITY DETERMINANT The role of intrinsic modality elements in OCR-2 was revealed from phenotypic analysis of the ocr-2(yz5) mutation, which is a single nucleotide change that results in a glycine-to-glutamate (G36E) substitution in the cytoplasmic region of the

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predicted OCR-2 protein (Zhang et al., 2004). The ocr-2(yz5) mutant was isolated from the genetic screen for serotonin synthesis mutants, and the mutant animals exhibited dramatic reduction of tph-1 expression in ADF, equivalent to the mutant carrying a deletion that removes the ﬁrst ﬁve transmembrane segments of OCR-2, or those carrying ocr-2/osm-9 double mutation. The ocr-2(yz5) mutants also show severe defects in osmotic sensation; however, they respond as well as wild-type animals to diacetyl sensation over the range of one-million-fold dilutions (Sokolchik et al., 2005). The OCR-2(G36E) substitution is located about 220 amino acids apart from the ﬁrst ankryin motif, and this region does not show strong homology to any conserved functional domains. Consistent with its normal function in diacetyl sensation, the OCR-2(G36E) protein appears to be expressed and properly localized in vivo (Figure 18.1). The substitution is unlikely to cause a global change in the assembly of the ankryin motifs, as two osm-9 alleles have different amino acid substitutions in the ankyrin motifs; both alleles affect every sensory function involving OCR-2/OSM-9. A clue to the mechanistic role of this mutation comes from expression of a chimeric channel of OCR-2 and another C. elegans TRPV channel protein OCR-4 in ocr-2 deletion mutants. OCR-4 itself cannot substitute for any OCR-2 function. Replacing the N-terminal segment preceding the ﬁrst ankyrin motif of OCR-2 for the cognate segment in OCR-4 signiﬁcantly rescued tph-1 expression and osmotic sensation, but the chimeric protein failed to rescue diacetyl sensation of ocr-2 deletion mutants, suggesting that the primary structure residing in the cytoplasmic N-terminus of OCR-2 governs a subset of sensory modalities of the channel. Because the glycine residue appears to be in a region conserved in OCR-4, and OCR-4 cannot substitute for OCR-2 function, the glycine itself may not be the modality determinant. One plausible function of this N-terminal region is gating the channel. Recent studies demonstrated that the antiparallel α-helices of ankyrin motifs stack to form a superhelical spiral, acting as a gate spring of TRPN1 channels in hair cells and in Drosophila bristles (Sotomayor et al., 2005; Lee et al., 2006). TRPN1 orthologues have dazzling 17 to 29 ankyrin repeats; the ankyrin spring is thought to directly sense mechanical force from sound waves or a tactile stimulus, thereby coupling the tension in the plane of the membrane bilayer to the gate of the channel (Walker et al., 2000; Sidi et al., 2003; Corey et al., 2004; Shin et al., 2005). However, atomic force microscopy showed that the most efﬁcient staking structure is generated from the proteins with four to six ankryin repeats (Lee et al., 2006), which is a common feature of the members of TRPV channel proteins, including OCR-2 (Tobin et al., 2002). In a hypothetical model, the N-terminal tip region of OCR-2 might fold into a moiety subserving as the door attached to the spring formed by the ankyrin repeats, and the G36E substitution alters the gating property (Figure 18.1). This model resembles the ‘‘ball-and-chain’’ gating used by the Shaker K+ channels, in which the N-terminal residues act as the ‘‘ball’’ to block the channel (Yellen, 2002). Consistent with the lack of sequence conservation among N-termini of TRPV channel proteins, there is little sequence homology between various N-termini capable of producing ‘‘ball’’ blockers in the Shaker K+ channel proteins. However, charges and hydrophobic characters of the “ball” are important for the interaction with the gate of the Shaker K+

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channel (Murrell-Lagnado and Aldrich, 1993). Intriguingly, the tyrosine-to-alanine substitution at the N-terminal part of the third transmembrane segment of mouse TRPV4 selectively impairs activation of TRPV4 by heat and phorbol esters but not by cell swelling or arachidonic acid (Vriens et al., 2004), and residues at the cytophasmic phase between the second and third transmembrane segments of mammalian TRPV1 confer the sensitivity to vanilloids (Jordt and Julius, 2002). It is possible that the N-terminal ‘‘ball’’ interacts with speciﬁc residues around the cavity of the TRPV channel, or the “ball” may recruit speciﬁc intracellular ligands to the gate, thereby coupling sensory information, channel activity, and the signaling cascades.

TRP CHANNELS IN MAMMALIAN SEROTONERGIC SYSTEMS? Identiﬁcation of OCR-2/OSM-9 as a regulator of serotonin production in C. elegans raises the question of whether TRP channels also function in mammalian serotonergic systems. There is not yet explicit evidence indicating that any mammalian TRPV protein is expressed in serotonergic neurons. However, there is compelling evidence that TRPV1–V5 are expressed broadly in the CNS, including the hippocampus, amygdala, and hypothalamus (Xu et al., 2002; Smith et al., 2002; Vennekens et al., 2002). These brain areas receive rich serotonergic innervation and are the commanders for sensory-endocrine integration, emotion, and cognitive functions. TRPV1 and V4 act as molecular sensors in vivo, and TRPV1–V4 are each activated by diverse chemical and physical stimuli in heterologous expression systems (Ramsey et al., 2006). Interestingly, intranigral injection of the TRPV1 ligand depleted serotonin level in the nigra (Dawbarn et al., 1981). Also, peripherally administered capsaicin resulted in changes in the electroencephalogram activity in the dorsal raphe, the area enriched with serotonergic nuclei (Rabe et al., 1980), and TRPV4 is a locus associated with bipolar affective disorders (Delany et al., 2001). These studies do not establish the TRPV1 and TRPV4 regulating serotonin signaling but they imply that TRPV channels play a broad role through their functions in the CNS and probably via the serotonergic system. Multiple sensory modalities, diverse gating mechanisms, and associations to distinct signaling cascades of OCR-2/OSM-9 provide insights into how a single channel entity can be dedicated to multiple sensory functions in vivo. The ﬁve mammalian TRPV channel genes expressed in the brain can produce a vast array of distinct channels through splicing variants, heteromeric combinations, and associations with distinct cellular components. These properties endow TRP channels to serve as molecular sensors in billions of years of metazoan evolution. It is tempting to speculate that these ancient molecular sensors and the ancient cellular censors, the serotonergic neurons, co-evolved. Like in C. elegans, TRP channels may function in a small subset of the serotonergic neurons in mammals. Alternatively, myriad diverse channels may underscore distinct serotonergic neurons in a wide spectrum of sensory functions. Even brain serotonergic neurons are not directly exposed to the external environment like the ADF neurons in C. elegans, but they may still sense chemical ligands and physical stimuli from other neuronal and nonneuronal cells and from the vesicular systems (Scheibel et al., 1975), therefore linking the

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gating information of TRP channels, sensory stimuli, serotonin production, and its downstream signaling.

REFERENCES Adell, A., Celada, P., Abellan, M.T., and Artigas, F. (2002). Origin and functional role of the extracellular serotonin in the midbrain raphe nuclei. Brain Res. Brain Res. Rev., 39:154–80. Review. Azmitia, E.C. (1999). Serotonin neurons, neuroplasticity, and homeostasis of neural tissue. Neuropsychopharmacology, 21:33S–45S. Review. Bailey, C.H., Chen, M., Keller, F., and Kandel, E.R. (1992). Serotonin-mediated endocytosis of apCAM: an early step of learning-related synaptic growth in Aplysia. Science, 256:645–49. Bargmann, C.I. and Horvitz, H.R. (1991). Chemosensory neurons with overlapping functions direct chemotaxis to multiple chemicals in C. elegans. Neuron, 7: 729–42. Berridge, M.J., Bootman, M.D., and Roderick, H.L. (2003). Calcium signalling: dynamics, homeostasis and remodelling. Nat. Rev. Mol. Cell Biol., 4:517–29. Review. Blier, P. and de Montigny, C. (1998). Possible serotonergic mechanisms underlying the antidepressant and anti-obsessive-compulsive disorder responses. Biological Psychiatry, 44:313–23. Boadle-Biber, M.C. (1993). Regulation of serotonin synthesis. Prog. Biophys. Mol. Biol., 60:1–15. Review. Brunelli, M., Castellucci, V., and Kandel, E.R. (1976). Synaptic facilitation and behavioral sensitization in Aplysia: possible role of serotonin and cyclic AMP. Science, 194:1178–81. Chaouloff, F., Berton, O., and Mormede, P. (1999). Serotonin and stress. Neuropsychopharmacology, 21:28S–32S. Review. Colbert, H.A. and Bargmann, C.I. (1997). Environmental signals modulate olfactory acuity, discrimination, and memory in Caenorhabditis elegans. Learn. Mem., 4:179–91. Colbert, H.A., Smith, T.L., and Bargmann, C.I. (1997). OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans. J. Neurosci., 17(21):8259–69. Corey, D.P., García-Añoveros, J., Holt, J.R., Kwan, K.Y., Lin, S.Y., Vollrath, M.A., Amalﬁtano, A., Cheung, E.L., Derﬂer, B.H., Duggan, A., Geleoc, G.S., Gray, P.A., Hoffman, M.P., Rehm, H.L., Tamasauskas, D., and Zhang, D.S. (2004). TRPA1 is a candidate for the mechanosensitive transduction channel of vertebrate hair cells. Nature, 432:723–30. Dawbarn, D., Harmar, A.J., and Pycock, C.J. (1981). Intranigral injection of capsaicin enhances motor activity and depletes nigral 5-hydroxytryptamine but not substance P. Neuropharmacology, 20:341–46. de Bono, M., Tobin, D.M., Davis, M.W., Avery, L., and Bargmann, C.I. (2002). Social feeding in Caenorhabditis elegans is induced by neurons that detect aversive stimuli. Nature, 419:899–903. Delany, N.S., Hurle, M., Facer, P., Alnadaf, T., Plumpton, C., Kinghorn, I., See, C.G., Costigan, M., Anand, P., Woolf, C.J., Crowther, D., Sanseau, P., and Tate, S.N. (2001). Identiﬁcation and characterization of a novel human vanilloid receptor-like protein, VRL2. Physiol. Genomics, 4:165–74. Dempsey, C.M., Mackenzie, S.M., Gargus, A., Blanco, G., and Sze, J.Y. (2005). Serotonin (5HT), ﬂuoxetine, imipramine and dopamine target distinct 5HT receptor signaling to modulate Caenorhabditis elegans egg-laying behavior. Genetics, 169:1425–36.

The TRPV Channel in C. elegans Serotonergic Neurons

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Desai, C., Garriga, G., McIntire, S.L., and Horvitz, H.R. (1988). A genetic pathway for the development of the Caenorhabditis elegans HSN motor neurons. Nature, 336:638–46. Duerr, J.S., Frisby, D.L., Gaskin, J., Duke, A., Asermely, K., Huddleston, D., Eiden, L.E., and Rand, J.B. (1999). The cat-1 gene of Caenorhabditis elegans encodes a vesicular monoamine transporter required for speciﬁc monoamine-dependent behaviors. J. Neurosci., 19:72–84. Hansson, S.R., Mezey, E., and Hoffman, B.J. (1998). Serotonin transporter messenger RNA in the developing rat brain: early expression in serotonergic neurons and transient expression in non-serotonergic neurons. Neuroscience, 83:1185–1201. Hare, E.E. and Loer, C.M. (2004). Function and evolution of the serotonin-synthetic bas-1 gene and other aromatic amino acid decarboxylase genes in Caenorhabditis. BMC Evol. Biol., 4:24. Hoffman, B.J., Hansson, S.R., Mezey, E., and Palkovits, M. (1998). Localization and dynamic regulation of biogenic amine transporters in the mammalian central nervous system. Frontiers in Neuroendocrinology, 19:187–231. Horvitz, H.R., Chalﬁe, M., Trent, C., Sulston, J.E., and Evans, P.D. (1982). 5HT and octopamine in the nematode C. elegans. Science, 216:1012–14. Hukema, R.K., Rademakers, S., Dekkers, M.P., Burghoorn, J., and Jansen, G. (2006). Antagonistic sensory cues generate gustatory plasticity in Caenorhabditis elegans. EMBO J., 25:312–22. Jordt, S.E. and Julius, D. (2002). Molecular basis for species-speciﬁc sensitivity to ‘‘hot’’ chili peppers. Cell, 108:421–30. Kahn-Kirby, A.H., Dantzker, J.L., Apicella, A.J., Schafer, W.R., Browse, J., Bargmann, C.I., and Watts, J.L. (2004). Speciﬁc polyunsaturated fatty acids drive TRPV-dependent sensory signaling in vivo. Cell, 119(6):889–900. Kandel, E.R. and Schwartz, J.H. (1982). Molecular biology of learning: modulation of transmitter release. Science, 218:433–43. Lebrand, C., Cases, O., Wehrle, R., Blakely, R.D., and Edwards, R.H. et al. (1998). Transient developmental expression of monoamine transporters in the rodent forebrain. Journal of Comparative Neurology, 401:506–24. Lee, G., Abdi, K., Jiang, Y., Michaely, P., Bennett, V., and Marszalek, P.E. (2006). Nanospring behaviour of ankyrin repeats. Nature, 440(7081):246–49. Leibowitz, S.F. and Alexander, J.T. (1998). Hypothalamic serotonin in control of eating behavior, meal size, and body weight. Biol. Psychiatry, 44:851–64. Review. Liedtke, W., Choe, Y., Marti-Renom, M.A., Bell, A.M., Denis, C.S., Sali, A., Hudspeth, A.J., Friedman, J.M., and Heller, S. (2000). Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor. Cell, 103:525–35. Liedtke, W., Tobin, D.M., Bargmann, C.I., and Friedman, J.M. (2003). Mammalian TRPV4 (VR-OAC) directs behavioral responses to osmotic and mechanical stimuli in Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA, 100:14531–36. Montell, C. (2005). The TRP superfamily of cation channels. Sci. STKE, Ref3. Murrell-Lagnado, R.D. and Aldrich, R.W. (1993). Interactions of amino terminal domains of Shaker K channels with a pore-blocking site studied with synthetic peptides. J. Gen. Physiol., 102:949–75. Nurrish, S., Segalat, L., and Kaplan, J.M. (1999). Serotonin inhibition of synaptic transmission: Galpha(0) decreases the abundance of UNC-13 at release sites. Neuron, 24:231–42. Pedersen, S.F., Owsianik, G., and Nilius, B. (2005). TRP channels: an overview. Cell Calcium, 38:233–52. Review. Rabe, L.S., Buck, S.H., Moreno, L., Burks, T.F., and Dafny, N. (1980). Neurophysiological and thermoregulatory effects of capsaicin. Brain Res. Bull., 5:755–58.

254

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Ramsey, I.S., Delling, M., and Clapham, D.E. (2006). An introduction to trp channels. Annu. Rev. Physiol., 68:619–47. Ranganathan, R., Cannon, S.C., and Horvitz, H.R. (2000). MOD-1 is a serotonin-gated chloride channel that modulates locomotory behaviour in C. elegans. Nature, 408:470–75. Ranganathan, R., Sawin, E.R., Trent, C., and Horvitz, H.R. (2001). Mutations in the Caenorhabditis elegans serotonin reuptake transporter MOD-5 reveal serotonindependent and -independent activities of ﬂuoxetine. J. Neurosci., 21:5871–84. Roayaie, K., Crump, J.G., Sagasti, A., and Bargmann, C.I. (1998). The G alpha protein ODR-3 mediates olfactory and nociceptive function and controls cilium morphogenesis in C. elegans olfactory neurons. Neuron, 20:55–67. Sawin, E.R., Ranganathan, R., and Horvitz, H.R. (2000). C. elegans locomotory rate is modulated by the environment through a dopaminergic pathway and by experience through a serotonergic pathway. Neuron, 26:619–31. Sengupta, P., Chou, J.H., and Bargmann, C.I. (1996). odr-10 encodes a seven transmembrane domain olfactory receptor required for responses to the odorant diacetyl. Cell, 84(6):899–909. Scheibel, M.E., Tomiyasu, U., and Scheibel, A.B. (1975). Do raphe nuclei of the reticular formation have a neurosecretory or vascular sensor function? Exp. Neurol., 47:316–29. Shin, J.B., Adams, D., Paukert, M., Siba, M., Sidi, S., Levin, M., Gillespie, P.G., and Grunder, S. (2005). Xenopus TRPN1 (NOMPC) localizes to microtubule-based cilia in epithelial cells, including inner-ear hair cells. Proc. Natl. Acad. Sci. USA, 102(35): 12572–77. Sidi, S., Friedrich, R.W., and Nicolson, T. (2003). NompC TRP channel required for vertebrate sensory hair cell mechanotransduction. Science, 301:96–99. Smith, G.D., Gunthorpe, M.J., Kelsell, R.E., Hayes, P.D., Reilly, P., Facer, P., Wright, J.E., Jerman, J.C., Walhin, J.P., Ooi, L., Egerton, J., Charles, K.J., Smart, D., Randall, A.D., Anand, P., and Davis, J.B. (2002). TRPV3 is a temperature-sensitive vanilloid receptor-like protein. Nature, 418:186–90. Sokolchik, I., Tanabe, T., Baldi, P.F., and Sze, J.Y. (2005). Polymodal sensory function of the Caenorhabditis elegans OCR-2 channel arises from distinct intrinsic determinants within the protein and is selectively conserved in mammalian TRPV proteins. J. Neurosci., 25:1015–23. Sotomayor, M., Corey, D.P., and Schulten, K. (2005). In search of the hair-cell gating spring elastic properties of ankyrin and cadherin repeats. Structure, 13:669–82. Sze, J.Y., Victor, M., Loer, C., Shi, Y., and Ruvkun, G. (2000). Food and metabolic signalling defects in a Caenorhabditis elegans serotonin-synthesis mutant. Nature, 403:560–64. Tobin, D., Madsen, D., Kahn-Kirby, A., Peckol, E., Moulder, G., Barstead, R., Maricq, A., and Bargmann, C. (2002). Combinatorial expression of TRPV channel proteins deﬁnes their sensory functions and subcellular localization in C. elegans neurons. Neuron, 35:307–18. Tsunoda, S., Sierralta, J., Sun, Y., Bodner, R., Suzuki, E., Becker, A., Socolich, M., and Zuker, C.S. (1997). A multivalent PDZ-domain protein assembles signalling complexes in a G-protein-coupled cascade. Nature, 388:243–49. Vennekens, R., Voets, T., Bindels, R.J., Droogmans, G., and Nilius, B. (2002). Current understanding of mammalian TRP homologues. Cell Calcium, 31:253–64. Review. Vriens, J., Watanabe, H., Janssens, A., Droogmans, G., Voets, T., and Nilius, B. (2004). Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4. Proc. Natl. Acad. Sci. USA, 101:396–401. Walker, R.G., Willingham, A.T., and Zuker, C.S. (2000). A Drosophila mechanosensory transduction channel. Science, 287:2229–34.

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Watanabe, H., Davis, J.B., Smart, D., Jerman, J.C., Smith, G.D., Hayes, P., Vriens, J., Cairns, W., Wissenbach, U., Prenen, J., Flockerzi, V., Droogmans, G., Benham, C.D., and Nilius, B. (2002). Activation of TRPV4 channels (hVRL-2/mTRP12) by phorbol derivatives. J. Biol. Chem., 277:13569–77. Weinshenker, D., Garriga, G., and Thomas, J.H. (1995). Genetic and pharmacological analysis of neurotransmitters controlling egg laying in C. elegans. J. Neurosci., 15:6975–85. White, J.G., Southgate, E., Thomson, J.N., and Brenner, S. (1986). The structure of the ventral nerve cord of Caenorhabditis elegans. Philos. Trans. R. Soc. Lond. B. Biol. Sci., 275:327–48. Xu, H., Ramsey, I.S., Kotecha, S.A., Moran, M.M., Chong, J.A., Lawson, D., Ge, P., Lilly, J., Silos-Santiago, I., Xie, Y., DiStefano, P.S., Curtis, R., and Clapham, D.E. (2002). TRPV3 is a calcium-permeable temperature-sensitive cation channel. Nature, 418:181–86. Yellen, G. (2005). The voltage-gated potassium channels and their relatives. Nature, 419(6902):35–42. Review. Zhang, Y., Lu, H., and Bargmann, C.I. (2005). Pathogenic bacteria induce aversive olfactory learning in Caenorhabditis elegans. Nature, 438:179–84. Zhang, S., Sokolchik, I., Blanco, G., and Sze, J.Y. (2004). Caenorhabditis elegans TRPV ion channel regulates 5HT biosynthesis in chemosensory neurons. Development, 131:1629–38.

19

TRP Channel Functioning in Mating and Fertilization X. Z. Shawn Xu University of Michigan

Maureen M. Barr University of Wisconsin at Madison

CONTENTS TRPP ......................................................................................................................257 Polycystin Ciliary Localization and ADPKD in C. elegans.....................257 The Kinesin KLP-6 Regulates TRPP Function and Localization ............259 Intraﬂagellar Transport Moves TRPV but Not TRPP in Cilia .................260 TRPP1 PLAT Binding Partners.................................................................260 Future Directions .......................................................................................261 The TRPC Channel Subfamily in C. elegans .......................................................262 TRP-3 Is Required for Sperm–Egg Interactions during Fertilization ................................................................................................262 TRP-3 Translocates from Intracellular Vesicles to the Plasma Membrane during Sperm Activation..................................263 Questions and Future Directions...............................................................264 Acknowledgments..................................................................................................264 References..............................................................................................................264

TRPP POLYCYSTIN CILIARY LOCALIZATION

AND

ADPKD

IN

C.

ELEGANS

Autosomal dominant polycystic kidney disease (ADPKD) is one of the most common monogenic diseases, affecting 1 in 400 to 1 in 1,000 individuals. In ADPKD patients, the kidney accumulates multiple cysts, which ultimately cause end-stage renal disease. Mutation in the PKD1 or PKD2 gene accounts for 95 percent of ADPKD cases.1,2 PKD1 encodes polycystin-1 (PC-1), a 4,302 amino acid protein with a large extracellular domain, a G-protein-coupled receptor proteolytic site (GPS), eleven transmembrane (TM) domains, and an intracellular C-terminus (Figure 19.1).1,3 The polycystin/lipoxygenase/alpha-toxin (PLAT) domain is located

257

258

TRP Ion Channel Function in Sensory Transduction

FIGURE 19.1 Polycystin localization on the ciliary membrane and interacting proteins. The KLP-6 kinesin-3 is required for polycystin-mediated sensory behaviors and TRPP localization. KLP-6 may act as a molecular motor transporting the TRPP complex on microtubules or as a molecular scaffold coupling the TRPP complex from the membrane to microtubules. The PLAT domain of LOV-1 physically associates with ATP-2 and KIN-10. ATP-2 is a component of the ATP synthase, localizes to the cilium, and regulates male mating behavior. KIN-10 is the regulatory subunit of casein kinase 2 (CK2). CK2 and TAX-6 calcineurin modulate PKD-2 ciliary localization and function. Adapted from reference 44.

in the ﬁrst cytoplasmic loop between TM1 and TM2 and has been postulated to be involved in membrane–protein or protein–protein interactions.4 The PLAT domain is conserved in all PC-1 family members and is also found in a variety of membraneor lipid-associated proteins. Polycystin-2 (PC-2, encoded by PKD2) is a transient receptor protein polycystin (TRPP) family member2 and acts as a nonselective cation channel (reviewed in reference 5). Mammalian PC-1 and PC-2 have been demonstrated to localize to primary cilia of kidney epithelial cells6,7 where they function as a mechanosensitive channel.8 ADPKD is one of a number of human genetic diseases that are rooted in defects in cilia formation, maintenance, or function.9,10,11 C. elegans provides a powerful model system to understand ADPKD and other diseases of ciliary basis.12 In C. elegans, LOV-1 and PKD-2 are homologous to polycystin-1 and polycystin-2, respectively.13,14,15 Similar to PC-1, LOV-1 has eleven predicted TM domains, an intracellular PLAT domain between TM1 and TM2, and a large extracellular domain. The extracellular domains of LOV-1 and PC-1 are divergent, suggesting that a species-speciﬁc ligand may activate the putative receptor LOV-1. LOV-1 does possess a GPS site, suggesting that proteolytic processing may occur as demonstrated for human PC-1 and sea urchin REJ3, a polycystin-like protein.16,17 Like PC-2, PKD-2 has six TM domains, an extracellular polycystin loop domain between TM1 and TM2, and a C-tail possessing coiled-coil and potential

TRP Channel Functioning in Mating and Fertilization

259

Ca2+-binding EF hand domains.14,18 LOV-1 and PKD-2 localize to ciliated endings on dendrites and in neuronal cell bodies of male-speciﬁc sensory neurons.13,14 lov-1 and pkd-2 are required for two mating behaviors (response to mate contact and location of the mate’s vulva) and are postulated to sense cues from the mate.13,14 Hence, the connection between the polycystins, cilia, and sensory function seems to be an ancient one. TRP vanilloid (TRPV) channels also localize to cilia in mammals, Drosophila, and the nematode.19,20,21 In C. elegans, the TRPV channels OSM-9 and OCR-2 depend on each other for ciliary localization and sensory function.22 In Drosophila, the ciliary localization of TRPV hearing channels Nanchung (NAN) and Inactive (IAV) is also codependent.23 Human PC-1 has been implicated in transporting PC-2 from the endoplasmic reticulum (ER) to the plasma membrane.24 In C. elegans sensory neurons, PKD-2 ciliary stabilization requires LOV-1, and vice versa.25 In a lov-1 mutant background, PKD-2 levels are greatly reduced in cilia. Hence, TRPP partnering is required for optimal ciliary targeting. Interestingly, PKD-2 forms abnormal aggregates in neuronal cell bodies of lov-1 mutants.25 Protein aggregation has not been examined or described in ADPKD cysts.

THE KINESIN KLP-6 REGULATES TRPP FUNCTION

AND

LOCALIZATION

The klp-6 mutant was identiﬁed based on its Rsp (response) and Lov (location of vulva) defects in the same genetic screen that yielded the lov-1/PC-1 mutant.26 In klp-6 mutants, PKD-2 abnormally accumulates at the base of cilia as opposed to the cilium proper and also accumulates along dendrites. klp-6 encodes a kinesin-like protein of 928 amino acids that belongs to the kinesin-3 (previously known as UNC104/Kif1A) family.27,28,29 Kinesin-3 family members are composed of an N-terminal, motor head region containing ATP- and microtubule-binding domains, followed by a coiled-coil and fork-head associated (FHA) domain that may facilitate multimerization,30 and a C-terminal tail of variable length and domain composition. Single homologues of KLP-6 are found in multiple vertebrate genomes. Alignments of C. elegans, zebraﬁsh, mouse, and rat KLP-6 revealed 34 percent identity and 52 percent similarity overall, a highly conserved motor domain (53 percent identity), and middle and tail domains conserved to a lesser extent. The C-terminal tail of kinesin enables many regulatory properties relating to cargo recognition, motor activity, and subcellular trafﬁcking.31–34 If the klp-6 mutant produces a protein, the truncated protein is predicted to be deﬁcient in cargo-binding but to retain microtubule-binding ability. KLP-6 is coexpressed and colocalized with LOV-1/PC-1 and PKD-2/PC-2 in malespeciﬁc sensory neurons. KLP-6 and the polycystins may function as an evolutionarily conserved ciliary unit (Figure 19.1). KLP-6 may act as a motor protein to directly transport multiple signaling molecules and distribute membrane receptors to speciﬁc ciliary subzones. Additionally, or possibly alternatively to the motor model, KLP-6 may function as a novel anchor-like protein and tether the polycystins and other unidentiﬁed proteins to ciliary microtubules. Mammalian polycystins function as mechanosensors in primary cilia of renal epithelial tubules, and mechanosensory complexes typically interact with microtubule or actin ﬁlaments via cytoskeletal-associated proteins.

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KLP-6 promises new routes to understanding cilia function, sensory behaviors, and potentially ADPKD.

INTRAFLAGELLAR TRANSPORT MOVES TRPV

BUT

NOT TRPP

IN

CILIA

The development of all cilia and ﬂagella requires intraﬂagellar transport (IFT).35 IFT is an evolutionarily conserved, microtubule-based motility ﬁrst observed as microscopic particles moving up and down the length of the ﬂagella of the green alga Chlamydomonas.36 The cellular machinery driving IFT was determined using primarily cellular and biochemical approaches.35 The IFT machinery contains heterotrimeric kinesin-2 and retrograde cytoplasmic dynein motors that move IFT particles and cargo to and from the distal tips of cilia. The IFT particle is composed of two complexes (A or B) containing 16 to 18 polypeptides. A simple model involves kinesin-2 and complex B polypeptides regulating anterograde transport, and dynein and complex A polypeptides regulating retrograde transport. In mammals, disruption of the kinesin-2 IFT motor or Polaris IFT complex B polypeptide results in embryos lacking cilia and exhibiting abnormal left-right development and polycystic kidney disease.37,38,39 Polycystin-2 is also required to establish left-right asymmetry in mice.40 PC-2 is found on nodal cilia and required for the generation of an asymmetric calcium signal, suggesting that PC-2 functions as a mechanosensor in nodal cilia.41 A role for IFT in transporting ciliary membrane proteins has recently been described. Using time-lapse ﬂuorescence microscopy and genetics, we demonstrated IFT-dependent vectorial transport of select GFP-tagged sensory receptors within the ciliary membrane of C. elegans sensory neurons in vivo.42 TRPV channels OSM-9 and OCR-221,22 move in cilia at rates comparable to the IFT machinery, and this motility is disrupted in IFT mutant backgrounds. Surprisingly, motility of TRPP channel PKD-2 is not detected, suggesting that PKD-2 may diffuse into the ciliary membrane, that PKD-2 may be physically restrained at the base of the cilium, or that at least two mechanisms regulate ciliary protein localization.

TRPP1 PLAT BINDING PARTNERS The evolutionarily conserved PLAT domain has been proposed to mediate protein– protein or protein–lipid interactions.4 In wild-type C. elegans males, overexpression of the PLAT domain alone or a LOV-1/PC-1 TM segment followed by the PLAT domain (TM-PLAT) interferes with male sensory behaviors.43 PLAT and TM-PLAT do not localize to cilia, indicating that the PLAT domain is not sufﬁcient for LOV1/PC-1 ciliary targeting.43 To identify targets of the C. elegans PLAT domain, a yeast two hybrid screen was performed.43 ATP-2, an ATP synthase subunit, and KIN-10, the regulatory beta subunit of the protein kinase CK2 (casein kinase 2), were isolated and validated.43,44 ATP-2 and KIN-10 also associate with the human PLAT domain. ATP-2 is a component of the ATP synthase, which is composed of two functional domains: a catalytic F1 portion and a membrane-embedded F0 portion. C. elegans ATP-2 is the beta subunit, or active site, of the F1 portion.45 The mitochondrial respiratory chain (MRC) generates the majority of cellular ATP. ATP-2 and other

TRP Channel Functioning in Mating and Fertilization

261

ATP synthase components but not MRC I–IV colocalize with PKD-2 in male-speciﬁc sensory cilia.43 Moreover, knockdown of the ATP synthase but not MRC I–IV produces polycystin-like male sensory defects.43 While the ATP synthase localizes primarily to the inner mitochondria membrane,46 its presence at the cell membrane has been reported.47–52 The ATP synthase has also been found in plasma membrane lipid rafts along with beta-tubulin.53,54 Our results suggest that the ciliary-localized ATP synthase may play a previously unsuspected role in polycystin signaling. C. elegans LOV-1 and human PC-1 also bind the regulatory subunit of CK2.44 Protein phosphorylation by the coordinated activities of protein kinases and phosphatases is central to many signal transduction pathways. CK2 and calcineurin/ protein phosphatase 2B (PP2B) modulate PKD-2 function and ciliary localization. CK2 and the Ca2+-activated phosphatase calcineurin act antagonistically to regulate PKD-2. A “phospho-defective” PKD-2 mutant protein trafﬁcks normally to cilia but exhibits attenuated function, while a “phospho-mimetic” PKD-2 is defective in both function and ciliary localization. tax-6 regulates PKD-2 ciliary localization but not ciliogenesis or gene expression. A dynamic phosphorylation cycle modulates normal polycystin function and ciliary distribution. Interestingly, CK2 has been implicated in the regulation of mammalian PC-2 activity and trafﬁcking to the plasma membrane,55,56 and PC-1 has been shown to activate a calcineurin/NFAT (nuclear factor of activated T-cells) signaling pathway.57 Mammalian PC-2S812 is constitutively phosphorylated in vivo.55 This CK2 site (S812) is not conserved in C. elegans PKD-2. Walz and colleagues have shown that trafﬁcking of PC-2 from the ER to the plasma membrane involves CK2 phosphorylation at S812 in an acidic cluster region and PACS proteins.56 There is no data to suggest that S812 is critical to PC-2 ciliary localization, and there is no acidic cluster region found in C. elegans PKD-2. Distinct mechanisms are likely required for localizing polycystin-2 to the plasma membrane and cilium. The PLAT domain of C. elegans LOV-1 and human PC-1 may coordinate CK2 localization and activity to a key PKD-2/PC-2 CK2 site (Figure 19.1). Upon stimulation, the polycystin mechanosensitive complex is activated8 and phosphorylated by CK2, causing an increase in intracellular Ca2+. Elevation in intracellular Ca2+ concentration activates TAX-6/calcineurin, resulting in dephosphorylation of PKD2/PC-2 and a return to an inactive state. Phosphorylation of PKD-2/PC-2 may serve as a mechanism for modulating channel properties; for attenuation of sensory signals; for clustering of channels, receptors, and signaling molecules; or for receptor internalization. In C. elegans, an imbalance between inactive and active PKD-2 states culminates in functional defects represented by a reduction in male mating behavior. CK2 and calcineurin have been individually implicated in other behaviors such as Drosophila circadian rhythms and mammalian learning and memory, respectively,58,59 as well as the regulation of ion channel activity or localization.55,56,60–64

FUTURE DIRECTIONS Recent genomic and proteomic approaches have identiﬁed components required for formation and function of cilia and ﬂagella.65–71 In contrast to ciliogenesis, very little is known regarding “sensorigenesis,” the process by which a cilium is specialized

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TRP Ion Channel Function in Sensory Transduction

for a particular function. Cilia often possess unique morphologies and express a distinct repertoire of sensory receptors and signaling molecules. The polycystins are required for the ﬂow-induced mechanosensory properties of kidney cilia,8,72,73,74 with defects resulting in ADPKD. How is this essential polycystin sensory complex regulated? How does the renal primary cilium sense urine ﬂow? A genome-wide expression proﬁling of ray-enriched genes identiﬁed novel cwp (coexpressed with polycystins) genes whose expression patterns are identical to pkd-2 and lov-1.75 These cell-type speciﬁc factors would regulate not only PKD-2 localization, but also contribute to the generation of sex-speciﬁc sensory behaviors. Future investigations of the genetic, cellular, molecular, and biochemical roles of these candidates and others in polycystin localization and function are warranted.

THE TRPC CHANNEL SUBFAMILY IN C. ELEGANS The C. elegans genome encodes three TRPC subfamily members: TRP-1, TRP-2, and TRP-3.15,76 TRP-1 and TRP-2 share 35–45 percent sequence identity with human TRPCs, while the homology between TRP-3 and human TRPCs is 25–30 percent. All three worm TRPCs possess the same domain structure as their mammalian counterparts. This includes three to four ankyrin repeats and a coiled-coil domain in the N-terminus, followed by six putative transmembrane domains and a TRP homology domain in the C-terminus. Although TRP-3 has been characterized in vivo (see below), functional analyses of TRP-1 and TRP-2 have not been reported.76

TRP-3 IS REQUIRED FOR SPERM–EGG INTERACTIONS DURING FERTILIZATION TRP-3 protein is enriched in sperm as evidenced by antibody staining.76 A microarray study also reveals that TRP-3 mRNA is enriched in sperm.77 Consistent with the expression pattern, trp-3 deletion mutants are nearly sterile with an average fertility of ~5 percent of wild-type hermaphrodites.76 This sterile phenotype is due to a defect in sperm, but not in oocytes or ovulation. Likewise, trp-3 males are also nearly sterile. Further analysis indicates that trp-3 mutant sperm are developmentally normal and motile and can make contact with oocytes in the spermatheca where sperm are stored and fertilization takes place. These results indicate that the sterile phenotype in trp-3 results from a defect in fertilization.76 Fertilization is triggered by a series of specialized sperm–egg interactions including gamete recognition, binding, and fusion; however, the molecular mechanisms underlying sperm–egg interactions are not well understood.78 Because of its facile genetics, C. elegans has recently emerged as a genetic model for the study of fertilization.79 Despite their morphological differences from mammalian sperm and lack of acrosome, nematode sperm have basic functions common to all sperm, including spermatogenesis, sperm activation (spermiogenesis), motility, gamete recognition/adhesion, and gamete fusion.79 In particular, the absence of egg coats in C. elegans oocytes greatly facilitates the analysis of fertilization.79 Thus, during nematode fertilization, gamete binding and fusion likely follow gamete contact/recognition. Interestingly, trp-3 mutant sperm can bind to the oocyte, suggesting that the mutant sperm may be defective in sperm–egg fusion.76 However, the lack of a robust

TRP Channel Functioning in Mating and Fertilization

263

in vitro sperm–egg binding assay in C. elegans precludes the measurement of sperm–egg binding afﬁnity. Thus, whether trp-3 mutations impair sperm–egg binding remains an open question. Notwithstanding, these analyses demonstrate that the sterile phenotype of trp-3 mutants results from a defect of trp-3 sperm in mediating sperm–egg interactions during fertilization. Are the functions of TRPC proteins conserved in fertilization? All seven mammalian TRPCs are found to be expressed in sperm and have been suggested to play roles in regulating sperm acrosome reaction and motility, but their roles in sperm–egg binding/fusion have not been examined.80,81 Speciﬁcally, mouse TRPC3 and TRPC6 are localized to the posterior part of the sperm head (the region that mediates sperm–egg binding and fusion), raising the possibility that these TRPCs might play roles in these processes.80,81 Nevertheless, genetic ablation of individual TRPCs in mice, such as TRPC2, TRPC4, and TRPC6, has not revealed a defect in fertility.82,83,84 It is possible that the functions of mouse TRPCs in fertilization are redundant.

TRP-3 TRANSLOCATES FROM INTRACELLULAR VESICLES MEMBRANE DURING SPERM ACTIVATION

TO THE

PLASMA

In spermatids, TRP-3 is localized to the membranous organelles (MOs), a class of intracellular vesicles derived from the ER/Golgi during spermatogenesis.76 By contrast, in mature sperm (spermatozoa), TRP-3 is localized to the plasma membrane.76 This suggests that TRP-3 undergoes protein translocation during sperm activation, a process by which spermatids develop into mature sperm. The MOs in spermatids fuse with the plasma membrane during sperm activation and become permanently attached to the plasma membranes of mature sperm. This process is also accompanied by the development of a pseudopod, resulting in the transformation of round immotile spermatids into polarized motile mature sperm (Figure 19.2). In the sterile fer-1 mutants, the MOs fail to fuse with the plasma membrane, while sperm activation proceeds.85 In support of the translocation model, TRP-3 no longer undergoes

laminar membrane

TRP-3

TRP-3

sperm activation

MO

pseudopod TRP-3 MO

spermatid

mature sperm

FIGURE 19.2 A schematic model showing TRP-3 translocation during sperm activation (spermiogenesis). The circles depict the MO vesicles, while the ovals represent TRP-3. Adapted from reference 76.

264

TRP Ion Channel Function in Sensory Transduction

protein translocation and is instead restricted to the MOs in fer-1 mutant sperm.76 Translocation of TRP-3 thus provides an in vivo mechanism for the regulation of TRP-3 function. A similar phenomenon has been observed with the Drosophila TRPC member TRPL and mammalian TRPC5 in response to light stimulation and epidermal growth factor treatment, respectively.86,87

QUESTIONS

AND

FUTURE DIRECTIONS

Questions remain as to how TRP-3 is activated in vivo and how TRP-3 activation leads to gamete fusion. Identifying the genes that genetically interact with trp-3 might provide insights into these questions. Several genes such as spe-9, spe-38, and spe-42, when mutated, lead to similar sperm defects to those of trp-3 mutants.88,89,90 These genes all encode membrane proteins, but their potential interactions with trp-3 have not been evaluated. As with many mammalian TRPCs, expression of TRP-3 in HEK293 cells promotes both receptor- and store-operated calcium entry.76 Because fertilization in nematodes occurs very rapidly after gamete contact,79 relatively slow kinetics of the store depletion makes it unlikely to be the physiological signal leading to TRP-3 activation in vivo. Thus, TRP-3 might function as a receptor-operated channel in sperm. If so, the binding of sperm receptors and oocyte ligands in the plasma membrane might signal the opening of TRP-3 in sperm, and the ensuing calcium inﬂux would then trigger a series of signaling events culminating in gamete fusion.

ACKNOWLEDGMENTS We thank Paul W. Sternberg (California Institute of Technology) for the intellectually challenging and rich scientiﬁc environment. Research in our laboratories is supported by the NIH (M.M.B. and X.Z.S.X.), PKD Foundation (M.M.B.), and University of Michigan BSSP Program (X.Z.S.X.).

REFERENCES 1. Hughes, J., Ward, C.J., Peral, B., Aspinwall, R., Clark, K., San Millan, J.L., Gamble, V. & Harris, P.C. (1995). The polycystic kidney disease 1 (PKD1) gene encodes a novel protein with multiple cell recognition domains. Nat. Genet. 10, 151–60. 2. Mochizuki, T., Wu, G., Hayashi, T., Xenophontos, S.L., Veldhuisen, B., Saris, J.J., Reynolds, D.M., Cai, Y., Gabow, P.A., Pierides, A., Kimberling, W.J., Breuning, M.H., Deltas, C.C., Peters, D.J. & Somlo, S. (1996). PKD2, a gene for polycystic kidney disease that encodes an integral membrane protein. Science 272, 1339–42. 3. Igarashi, P. & Somlo, S. (2002). Genetics and pathogenesis of polycystic kidney disease. J. Am. Soc. Nephrol. 13, 2384–98. 4. Bateman, A. & Sandford, R. (1999). The PLAT domain: a new piece in the PKD1 puzzle. Curr. Biol. 9, R588–90. 5. Corey, D.P. (2003). New TRP channels in hearing and mechanosensation. Neuron 39, 585–88.

TRP Channel Functioning in Mating and Fertilization

265

6. Pazour, G.J., San Agustin, J.T., Follit, J.A., Rosenbaum, J.L. & Witman, G.B. (2002). Polycystin-2 localizes to kidney cilia and the ciliary level is elevated in orpk mice with polycystic kidney disease. Curr. Biol. 12, R378–80. 7. Yoder, B.K., Hou, X. & Guay-Woodford, L.M. (2002). The polycystic kidney disease proteins, polycystin-1, polycystin-2, polaris, and cystin, are co-localized in renal cilia. J. Am. Soc. Nephrol. 13, 2508–16. 8. Nauli, S.M., Alenghat, F.J., Luo, Y., Williams, E., Vassilev, P., Li, X., Elia, A.E., Lu, W., Brown, E.M., Quinn, S.J., Ingber, D.E. & Zhou, J. (2003). Polycystins 1 and 2 mediate mechanosensation in the primary cilium of kidney cells. Nat. Genet. 33, 129–37. 9. Pazour, G.J. & Rosenbaum, J.L. (2002). Intraﬂagellar transport and cilia-dependent diseases. Trends Cell Biol. 12, 551–55. 10. Watnick, T. & Germino, G. (2003). From cilia to cyst. Nat. Genet. 34, 355–56. 11. Pazour, G.J. (2004). Intraﬂagellar transport and cilia-dependent renal disease: the ciliary hypothesis of polycystic kidney disease. J. Am. Soc. Nephrol. 15, 2528–36. 12. Barr, M.M. (2005). Caenorhabditis elegans as a model to study renal development and disease: sexy cilia. J. Am. Soc. Nephrol. 16, 305–12. 13. Barr, M.M. & Sternberg, P.W. (1999). A polycystic kidney-disease gene homologue required for male mating behaviour in C. elegans. Nature 401, 386–89. 14. Barr, M.M., DeModena, J., Braun, D., Nguyen, C.Q., Hall, D.H. & Sternberg, P.W. (2001). The Caenorhabditis elegans autosomal dominant polycystic kidney disease gene homologs lov-1 and pkd-2 act in the same pathway. Curr. Biol. 11, 1341–46. 15. Kahn-Kirby, A.H. & Bargmann, C.I. (2006). TRP channels in C. elegans. Annu. Rev. Physiol. 68, 719–36. 16. Qian, F., Boletta, A., Bhunia, A.K., Xu, H., Liu, L., Ahrabi, A.K., Watnick, T.J., Zhou, F. & Germino, G.G. (2002). Cleavage of polycystin-1 requires the receptor for egg jelly domain and is disrupted by human autosomal-dominant polycystic kidney disease 1–associated mutations.Proc. Natl. Acad. Sci. USA 99, 16981–86. 17. Mengerink, K.J., Moy, G.W. & Vacquier, V.D. (2002). suREJ3, a polycystin-1 protein, is cleaved at the GPS domain and localizes to the acrosomal region of sea urchin sperm. J. Biol. Chem. 277, 943–48. 18. Koulen, P., Duncan, R.S., Liu, J., Cohen, N.E., Yannazzo, J.A., McClung, N., Lockhart, C.L., Branden, M. & Buechner, M. (2005). Polycystin-2 accelerates Ca2+ release from intracellular stores in Caenorhabditis elegans. Cell Calcium 37, 593–601. 19. Andrade, Y.N., Fernandes, J., Vazquez, E., Fernandez-Fernandez, J.M., Arniges, M., Sanchez, T.M., Villalon, M. & Valverde, M.A. (2005). TRPV4 channel is involved in the coupling of ﬂuid viscosity changes to epithelial ciliary activity. J. Cell Biol. 168, 869–74. 20. Kim, J., Chung, Y.D., Park, D.Y., Choi, S., Shin, D.W., Soh, H., Lee, H.W., Son, W., Yim, J., Park, C.S., Kernan, M.J. & Kim, C. (2003). A TRPV family ion channel required for hearing in Drosophila. Nature 424, 81–84. 21. Colbert, H.A., Smith, T.L. & Bargmann, C.I. (1997). OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans. J. Neurosci. 17, 8259–69. 22. Tobin, D., Madsen, D., Kahn-Kirby, A., Peckol, E., Moulder, G., Barstead, R., Maricq, A. & Bargmann, C. (2002). Combinatorial expression of TRPV channel proteins deﬁnes their sensory functions and subcellular localization in C. elegans neurons. Neuron 35, 307–18.

266

TRP Ion Channel Function in Sensory Transduction

23. Gong, Z., Son, W., Chung, Y.D., Kim, J., Shin, D.W., McClung, C.A., Lee, Y., Lee, H.W., Chang, D.J., Kaang, B.K., Cho, H., Oh, U., Hirsh, J., Kernan, M.J. & Kim, C. (2004). Two interdependent TRPV channel subunits, inactive and Nanchung, mediate hearing in Drosophila. J. Neurosci. 24, 9059–66. 24. Hanaoka, K., Qian, F., Boletta, A., Bhunia, A.K., Piontek, K., Tsiokas, L., Sukhatme, V.P., Guggino, W.B. & Germino, G.G. (2000). Co-assembly of polycystin-1 and -2 produces unique cation-permeable currents. Nature 408, 990–94. 25. Bae, Y.K., Qin, H., Knobel, K.M., Hu, J., Rosenbaum, J.L. & Barr, M.M. (2006). General and cell-type speciﬁc mechanisms target TRPP2 to cilia PKD-2. Development, in press. 26. Peden, E.M. & Barr, M.M. (2005). The KLP-6 kinesin is required for male mating behaviors and polycystin localization in Caenorhabditis elegans. Curr. Biol. 15, 394–404. 27. Siddiqui, S.S. (2002). Metazoan motor models: kinesin superfamily in C. elegans. Trafﬁc 3, 20–28. 28. Vale, R.D. (2003). The molecular motor toolbox for intracellular transport. Cell 112, 467–80. 29. Lawrence, C.J., Dawe, R.K., Christie, K.R., Cleveland, D.W., Dawson, S.C., Endow, S.A., Goldstein, L.S., Goodson, H.V., Hirokawa, N., Howard, J., Malmberg, R.L., McIntosh, J.R., Miki, H., Mitchison, T.J., Okada, Y., Reddy, A.S., Saxton, W.M., Schliwa, M., Scholey, J.M., Vale, R.D., Walczak, C.E. & Wordeman, L. (2004). A standardized kinesin nomenclature. J. Cell Biol. 167, 19–22. 30. Lee, J.R., Shin, H., Choi, J., Ko, J., Kim, S., Lee, H.W., Kim, K., Rho, S.H., Lee, J.H., Song, H.E., Eom, S.H. & Kim, E. (2004). An intramolecular interaction between the FHA domain and a coiled coil negatively regulates the kinesin motor KIF1A. Embo J. 23, 1506–15. 31. Nakagawa, T., Setou, M., Seog, D., Ogasawara, K., Dohmae, N., Takio, K. & Hirokawa, N. (2000). A novel motor, KIF13A, transports mannose-6-phosphate receptor to plasma membrane through direct interaction with AP-1 complex. Cell 103, 569–81. 32. Coy, D.L., Hancock, W.O., Wagenbach, M. & Howard, J. (1999). Kinesin’s tail domain is an inhibitory regulator of the motor domain. Nat. Cell Biol. 1, 288–92. 33. Setou, M., Seog, D.H., Tanaka, Y., Kanai, Y., Takei, Y., Kawagishi, M. & Hirokawa, N. (2002). Glutamate-receptor-interacting protein GRIP1 directly steers kinesin to dendrites. Nature 417, 83–87. 34. Seiler, S., Kirchner, J., Horn, C., Kallipolitou, A., Woehlke, G. & Schliwa, M. (2000). Cargo binding and regulatory sites in the tail of fungal conventional kinesin. Nat. Cell Biol. 2, 333–38. 35. Rosenbaum, J.L. & Witman, G.B. (2002). Intraﬂagellar transport. Nat. Rev. Mol. Cell Biol. 3, 813–25. 36. Kozminski, K.G., Forscher, P. & Rosenbaum, J.L. (1998). Three ﬂagellar motilities in Chlamydomonas unrelated to ﬂagellar beating. Video supplement. Cell Motil. Cytoskeleton 39, 347–48. 37. Lin, F., Hiesberger, T., Cordes, K., Sinclair, A.M., Goldstein, L.S., Somlo, S. & Igarashi, P. (2003). Kidney-speciﬁc inactivation of the KIF3A subunit of kinesin-II inhibits renal ciliogenesis and produces polycystic kidney disease. Proc. Natl. Acad. Sci. USA. 38. Murcia, N.S., Richards, W.G., Yoder, B.K., Mucenski, M.L., Dunlap, J.R. & Woychik, R.P. (2000). The Oak Ridge polycystic kidney (orpk) disease gene is required for left-right axis determination. Development 127, 2347–55.

TRP Channel Functioning in Mating and Fertilization

267

39. Taulman, P.D., Haycraft, C.J., Balkovetz, D.F. & Yoder, B.K. (2001). Polaris, a protein involved in left-right axis patterning, localizes to basal bodies and cilia. Mol. Biol. Cell 12, 589–99. 40. Wu, G., Markowitz, G.S., Li, L., D’Agati, V.D., Factor, S.M., Geng, L., Tibara, S., Tuchman, J., Cai, Y., Park, J.H., van Adelsberg, J., Hou, H., Jr., Kucherlapati, R., Edelmann, W. & Somlo, S. (2000). Cardiac defects and renal failure in mice with targeted mutations in Pkd2. Nat. Genet. 24, 75–78. 41. McGrath, J., Somlo, S., Makova, S., Tian, X. & Brueckner, M. (2003). Two populations of node monocilia initiate left-right asymmetry in the mouse. Cell 114, 61–73. 42. Qin, H., Burnette, D.T., Bae, Y.K., Forscher, P., Barr, M.M. & Rosenbaum, J.L. (2005). Intraﬂagellar transport is required for the vectorial movement of TRPV channels in the ciliary membrane. Curr. Biol. 15, 1695–99. 43. Hu, J. & Barr, M.M. (2005). ATP-2 interacts with the PLAT domain of LOV-1 and is involved in Caenorhabditis elegans polycystin signaling. Mol. Biol. Cell 16, 458–69. 44. Hu, J., Bae, Y.-K., Knobel, K.M. & Barr, M.M. (2006). Casein kinase II and calcineurin modulate TRPP function and ciliary localization. Mol. Biol. Cell. 17, 2200–11. 45. Tsang, W.Y., Sayles, L.C., Grad, L.I., Pilgrim, D.B. & Lemire, B.D. (2001). Mitochondrial respiratory chain deﬁciency in Caenorhabditis elegans results in developmental arrest and increased life span. J. Biol. Chem. 276, 32240–46. 46. Boyer, P.D. (1997). The ATP synthase—a splendid molecular machine. Annu. Rev. Biochem. 66, 717–49. 47. Das, B., Mondragon, M.O., Sadeghian, M., Hatcher, V.B. & Norin, A.J. (1994). A novel ligand in lymphocyte-mediated cytotoxicity: expression of the beta subunit of H+ transporting ATP synthase on the surface of tumor cell lines. J. Exp. Med. 180, 273–81. 48. Moser, T.L., Kenan, D.J., Ashley, T.A., Roy, J.A., Goodman, M.D., Misra, U.K., Cheek, D.J. & Pizzo, S.V. (2001). Endothelial cell surface F1-F0 ATP synthase is active in ATP synthesis and is inhibited by angiostatin. Proc. Natl. Acad. Sci. USA 98, 6656–61. 49. Moser, T.L., Stack, M.S., Asplin, I., Enghild, J.J., Hojrup, P., Everitt, L., Hubchak, S., Schnaper, H.W. & Pizzo, S.V. (1999). Angiostatin binds ATP synthase on the surface of human endothelial cells. Proc. Natl. Acad. Sci. USA 96, 2811–16. 50. Chang, S.Y., Park, S.G., Kim, S. & Kang, C.Y. (2002). Interaction of the C-terminal domain of p43 and the alpha subunit of ATP synthase. Its functional implication in endothelial cell proliferation. J. Biol. Chem. 277, 8388–94. 51. Arakaki, N., Nagao, T., Niki, R., Toyofuku, A., Tanaka, H., Kuramoto, Y., Emoto, Y., Shibata, H., Magota, K. & Higuti, T. (2003). Possible role of cell surface H(+)-ATP synthase in the extracellular ATP synthesis and proliferation of human umbilical vein endothelial cells. Mol. Cancer Res. 1, 931–39. 52. Martinez, L.O., Jacquet, S., Esteve, J.P., Rolland, C., Cabezon, E., Champagne, E., Pineau, T., Georgeaud, V., Walker, J.E., Terce, F., Collet, X., Perret, B. & Barbaras, R. (2003). Ectopic beta-chain of ATP synthase is an apolipoprotein A-I receptor in hepatic HDL endocytosis. Nature 421, 75–79. 53. Bae, T.J., Kim, M.S., Kim, J.W., Kim, B.W., Choo, H.J., Lee, J.W., Kim, K.B., Lee, C.S., Kim, J.H., Chang, S.Y., Kang, C.Y., Lee, S.W. & Ko, Y.G. (2004). Lipid raft proteome reveals ATP synthase complex in the cell surface. Proteomics 4, 3536–48.

268

TRP Ion Channel Function in Sensory Transduction

54. Li, N., Shaw, A.R., Zhang, N., Mak, A. & Li, L. (2004). Lipid raft proteomics: Analysis of in-solution digest of sodium dodecyl sulfate-solubilized lipid raft proteins by liquid chromatography-matrix-assisted laser desorption/ionization tandem mass spectrometry. Proteomics 4, 3156–66. 55. Cai, Y., Anyatonwu, G., Okuhara, D., Lee, K.B., Yu, Z., Onoe, T., Mei, C.L., Qian, Q., Geng, L., Wiztgall, R., Ehrlich, B.E. & Somlo, S. (2004). Calcium dependence of polycystin-2 channel activity is modulated by phosphorylation at Ser812. J. Biol. Chem. 279, 19987–95. 56. Kottgen, M., Benzing, T., Simmen, T., Tauber, R., Buchholz, B., Feliciangeli, S., Huber, T.B., Schermer, B., Kramer-Zucker, A., Hopker, K., Simmen, K.C., Tschucke, C.C., Sandford, R., Kim, E., Thomas, G. & Walz, G. (2005). Trafﬁcking of TRPP2 by PACS proteins represents a novel mechanism of ion channel regulation. Embo J. 24, 705–16. 57. Puri, S., Magenheimer, B.S., Maser, R.L., Ryan, E.M., Zien, C.A., Walker, D.D., Wallace, D.P., Hempson, S.J. & Calvet, J.P. (2004). Polycystin-1 activates the calcineurin/NFAT (nuclear factor of activated T-cells) signaling pathway. J. Biol. Chem. 279, 55455–64. 58. Blau, J. (2003). A new role for an old kinase: CK2 and the circadian clock. Nat. Neurosci. 6, 208–10. 59. Lee, J.I. & Ahnn, J. (2004). Calcineurin in animal behavior. Mol. Cells 17, 390–96. 60. Kuhara, A., Inada, H., Katsura, I. & Mori, I. (2002). Negative regulation and gain control of sensory neurons by the C. elegans calcineurin TAX-6. Neuron 33, 751–63. 61. Misonou, H., Mohapatra, D.P., Park, E.W., Leung, V., Zhen, D., Misonou, K., Anderson, A.E. & Trimmer, J.S. (2004). Regulation of ion channel localization and phosphorylation by neuronal activity. Nat. Neurosci. 7, 711–18. 62. Wu, Z.Z., Chen, S.R. & Pan, H.L. (2005). TRPV1 activation downregulates voltagegated calcium channels through calcium-dependant calcineurin in sensory neurons. J. Biol. Chem. 280, 18142–51. 63. Mohapatra, D.P. & Nau, C. (2005). Regulation of Ca2+-dependent desensitization in the vanilloid receptor TRPV1 by calcineurin and cAMP-dependent protein kinase. J. Biol. Chem. 280, 13424–32. 64. Bildl, W., Strassmaier, T., Thurm, H., Andersen, J., Eble, S., Oliver, D., Knipper, M., Mann, M., Schulte, U., Adelman, J.P. & Fakler, B. (2004). Protein kinase CK2 is coassembled with small conductance Ca(2+)-activated K+ channels and regulates channel gating. Neuron 43, 847–58. 65. Ostrowski, L.E., Blackburn, K., Radde, K.M., Moyer, M.B., Schlatzer, D.M., Moseley, A. & Boucher, R.C. (2002). A proteomic analysis of human cilia: identiﬁcation of novel components. Mol. Cell Proteomics 1, 451–65. 66. Avidor-Reiss, T., Maer, A.M., Koundakjian, E., Polyanovsky, A., Keil, T., Subramaniam, S. & Zuker, C.S. (2004). Decoding cilia function: deﬁning specialized genes required for compartmentalized cilia biogenesis. Cell 117, 527–39. 67. Blacque, O.E., Perens, E.A., Boroevich, K.A., Inglis, P.N., Li, C., Warner, A., Khattra, J., Holt, R.A., Ou, G., Mah, A.K., McKay, S.J., Huang, P., Swoboda, P., Jones, S.J., Marra, M.A., Baillie, D.L., Moerman, D.G., Shaham, S. & Leroux, M.R. (2005). Functional genomics of the cilium, a sensory organelle. Curr. Biol. 15, 935–41. 68. Eﬁmenko, E., Bubb, K., Mak, H.Y., Holzman, T., Leroux, M.R., Ruvkun, G., Thomas, J.H. & Swoboda, P. (2005). Analysis of xbx genes in C. elegans. Development 132, 1923–34.

TRP Channel Functioning in Mating and Fertilization

269

69. Li, J.B., Gerdes, J.M., Haycraft, C.J., Fan, Y., Teslovich, T.M., May-Simera, H., Li, H., Blacque, O.E., Li, L., Leitch, C.C., Lewis, R.A., Green, J.S., Parfrey, P.S., Leroux, M.R., Davidson, W.S., Beales, P.L., Guay-Woodford, L.M., Yoder, B.K., Stormo, G.D., Katsanis, N. & Dutcher, S.K. (2004). Comparative genomics identiﬁes a ﬂagellar and basal body proteome that includes the BBS5 human disease gene. Cell 117, 541–52. 70. Keller, L.C., Romijn, E.P., Zamora, I., Yates, J.R., 3rd & Marshall, W.F. (2005). Proteomic analysis of isolated chlamydomonas centrioles reveals orthologs of ciliarydisease genes. Curr. Biol. 15, 1090–98. 71. Pazour, G.J., Agrin, N., Leszyk, J. & Witman, G.B. (2005). Proteomic analysis of a eukaryotic cilium. J. Cell Biol. 170, 103–13. 72. Praetorius, H.A. & Spring, K.R. (2001). Bending the MDCK cell primary cilium increases intracellular calcium. J. Membr. Biol. 184, 71–79. 73. Praetorius, H.A. & Spring, K.R. (2003). Removal of the MDCK cell primary cilium abolishes ﬂow sensing. J. Membr. Biol. 191, 69–76. 74. Praetorius, H.A. & Spring, K.R. (2003). The renal cell primary cilium functions as a ﬂow sensor. Curr. Opin. Nephrol. Hypertens. 12, 517–20. 75. Portman, D.S. & Emmons, S.W. (2004). Identiﬁcation of C. elegans sensory ray genes using whole-genome expression proﬁling. Dev. Biol. 270, 499–512. 76. Xu, X.Z. & Sternberg, P.W. (2003). A C. elegans sperm TRP protein required for sperm-egg interactions during fertilization. Cell 114, 285–97. 77. Reinke, V., Smith, H.E., Nance, J., Wang, J., Van Doren, C., Begley, R., Jones, S.J., Davis, E.B., Scherer, S., Ward, S. & Kim, S.K. (2000). A global proﬁle of germline gene expression in C. elegans. Mol. Cell 6, 605–16. 78. Jungnickel, M.K., Sutton, K.A. & Florman, H.M. (2003). In the beginning: lessons from fertilization in mice and worms. Cell 114, 401–14. 79. Geldziler, B., Kadandale, P. & Singson, A. (2004). Molecular genetic approaches to studying fertilization in model systems. Reproduction 127, 409–16. 80. Sutton, K.A., Jungnickel, M.K., Wang, Y., Cullen, K., Lambert, S. & Florman, H.M. (2004). Enkurin is a novel calmodulin and TRPC channel binding protein in sperm. Dev. Biol. 274, 426–35. 81. Castellano, L.E., Trevino, C.L., Rodriguez, D., Serrano, C.J., Pacheco, J., Tsutsumi, V., Felix, R. & Darszon, A. (2003). Transient receptor potential (TRPC) channels in human sperm: expression, cellular localization and involvement in the regulation of ﬂagellar motility. FEBS Lett. 541, 69–74. 82. Freichel, M., Suh, S.H., Pfeifer, A., Schweig, U., Trost, C., Weissgerber, P., Biel, M., Philipp, S., Freise, D., Droogmans, G., Hofmann, F., Flockerzi, V. & Nilius, B. (2001). Lack of an endothelial store-operated Ca2+ current impairs agonist-dependent vasorelaxation in TRP4-/- mice. Nat. Cell Biol. 3, 121–27. 83. Leypold, B.G., Yu, C.R., Leinders-Zufall, T., Kim, M.M., Zufall, F. & Axel, R. (2002). Altered sexual and social behaviors in trp2 mutant mice. Proc. Natl. Acad. Sci. USA 99, 6376–81. 84. Stowers, L., Holy, T.E., Meister, M., Dulac, C. & Koentges, G. (2002). Loss of sex discrimination and male-male aggression in mice deﬁcient for TRP2. Science 295, 1493–500. 85. Achanzar, W.E. & Ward, S. (1997). A nematode gene required for sperm vesicle fusion. J. Cell Sci. 110 (Pt. 9), 1073–81. 86. Bahner, M., Frechter, S., Da Silva, N., Minke, B., Paulsen, R. & Huber, A. (2002). Light-regulated subcellular translocation of Drosophila TRPL channels induces longterm adaptation and modiﬁes the light-induced current. Neuron 34, 83–93.

270

TRP Ion Channel Function in Sensory Transduction

87. Bezzerides, V.J., Ramsey, I.S., Kotecha, S., Greka, A. & Clapham, D.E. (2004). Rapid vesicular translocation and insertion of TRP channels. Nat. Cell Biol. 6, 709–20. 88. Singson, A., Mercer, K.B. & L’Hernault, S.W. (1998). The C. elegans spe-9 gene encodes a sperm transmembrane protein that contains EGF-like repeats and is required for fertilization. Cell 93, 71–79. 89. Chatterjee, I., Richmond, A., Putiri, E., Shakes, D.C. & Singson, A. (2005). The Caenorhabditis elegans spe-38 gene encodes a novel four-pass integral membrane protein required for sperm function at fertilization. Development 132, 2795–808. 90. Kroft, T.L., Gleason, E.J. & L’Hernault, S.W. (2005). The spe-42 gene is required for sperm-egg interactions during C. elegans fertilization and encodes a spermspeciﬁc transmembrane protein. Dev. Biol. 286, 169–81.

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The Role of TRP Channels in Thermosensation Makoto Tominaga National Institutes of Natural Sciences

CONTENTS Introduction............................................................................................................271 Heat Receptors.......................................................................................................273 TRPV1 (VR1)............................................................................................273 TRPV2 (VRL-1) ........................................................................................275 Warm Receptors.....................................................................................................276 TRPV3 and TRPV4...................................................................................276 TRPM4 and TRPM5..................................................................................277 Cold Receptors ......................................................................................................278 TRPM8 (CMR1)........................................................................................278 TRPA1 (ANKTM1) ...................................................................................279 Thermal Nociception through TRP Channels in Invertebrates.............................280 Conclusion .............................................................................................................281 References..............................................................................................................281

INTRODUCTION We feel a wide range of temperatures spanning from coldness to heat. Within this range, temperatures over about 43°C and below about 15°C evoke not only a thermal sensation, but also a feeling of pain (LaMotte and Campbell, 1978; Tillman et al., 1995). Neurophysiological studies have demonstrated that the heat threshold of socalled C-ﬁber mechanoheat nociceptors (CMHs) depends on the absolute temperature, rather than the rate of temperature increase, and that the transduction of heat stimuli occurs at different skin depths for different CMHs (Tillman et al., 1995). Extreme cold also activates a subset of nociceptive neurons. However, the physiology of coldevoked pain is not as well understood as that of heat-evoked pain. It has been hypothesized that cutaneous nociceptor endings detect temperature and other physical stimuli by means of ion channels responsive to these stimuli. The ﬁrst support for this hypothesis came from the identiﬁcation of heat-gated ion channels present in a subset of primary afferent neurons (Cesare and McNaughton, 1996; Reichling and Levine, 1997). Insight into the molecular nature of these channels came shortly thereafter, with the cloning of the capsaicin receptor, TRPV1 (also known as VR1, 271

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the ﬁrst member of the TRPV subfamily) and the recognition that this ion channel protein, like mammalian nociceptors, could be activated by elevated temperatures with a discrete threshold near 43°C (Caterina et al., 1997; Caterina and Julius, 2001). Three other TRPV channels—TRPV2 (also known as VRL-1), TRPV3, and TRPV4 (also known as VROAC or OTRPC4)—have been cloned and characterized as heat or warm thermosensors (Jordt et al., 2003; Patapoutian et al., 2003; Tominaga and Caterina, 2004). In addition, two TRPM channels (TRPM4 and TRPM5) have been recently reported to be thermosensitive (Talavera et al., 2005). The threshold temperatures for activation of these channels range from relatively warm (TRPV3, TRPV4, TRPM4, and TRPM5) to extremely hot (TRPV2). In contrast to these warmth- or heat-activated TRP channels, two other TRP channels—TRPM8 (also known as CMR1) and TRPA1 (also known as ANKTM1)—are activated by cold stimuli (Jordt et al., 2003; Patapoutian et al., 2003; Tominaga and Caterina, 2004). This chapter focuses on eight mammalian thermosensitive TRP channels (Figure 20.1).

FIGURE 20.1 (Color ﬁgure follows p. 234.) (A) Temperature ranges activating thermosensitive TRP channels. (B) Phylogenetic relationship among the mammalian TRPV, TRPM, and TRPA channels with two Drosophila TRPA channels. Red, orange, and blue squares indicate channels activated by high heat, warm stimuli, and cold stimuli, respectively.

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HEAT RECEPTORS TRPV1 (VR1) When a receptor for capsaicin, a main pungent ingredient of hot chili peppers, was isolated using a Ca2+-imaging–based expression cloning method in 1997, it was designated vanilloid receptor subtype 1 (VR1), then renamed as TRPV1 (Caterina et al., 1997; Caterina and Julius, 2001). Patch-clamp recordings from human embryonic kidney-derived HEK293 cells transfected with TRPV1 cDNA (Caterina et al., 1997; Tominaga et al., 1998) revealed that capsaicin-activated TRPV1 currents exhibit nonselective cation permeability with an outwardly-rectifying currentvoltage (I-V) relationship. TRPV1 was also found to show high Ca2+ permeability (PCa/PNa= 9.6), which may be one of the reasons why the Ca2+-imaging strategy successfully cloned this molecule. Application of capsaicin to membrane patches excised from HEK293 cells expressing TRPV1 was shown to evoke clear singlechannel openings (conductance of ∼77 pS for Na+), strongly suggesting that no cytosolic second messengers are necessary for TRPV1 activation. TRPV1 transcript and protein were found to be most highly expressed in sensory neurons, especially in small diameter neurons within dorsal root and trigeminal sensory ganglia (DRG and TG, respectively), probably cell bodies of unmyelinated C-ﬁbers (Caterina et al., 1997; Tominaga et al., 1998), consistent with the notion that polymodal C-ﬁbers are involved in nociception. The burning quality of capsaicininduced pain suggests that capsaicin and heat may evoke painful responses through a common molecular pathway. Consistent with this hypothesis, TRPV1 was found to be activated by heat at >43°C, a temperature threshold similar to that at which heat evokes pain in vivo (Caterina et al., 1997; Tominaga et al., 1998). Single-channel openings recorded in excised-membrane patches expressing TRPV1 suggested that heat gates TRPV1 directly and that TRPV1 is itself a heat sensor. These ﬁndings also suggested that TRPV1 might constitute the heat-activated ion channels that have been characterized electrophysiologically in cultured sensory neurons and found to be sensitized through the protein kinase C-ε dependent pathway (Cesare et al., 1999). A strong correlation between nociceptor capsaicin responsiveness and heat responsiveness (Kirschstein et al., 1997; Nagy and Rang, 1999) provided further support for this idea. Additional studies demonstrated that TRPV1 could alternatively be activated at room temperature when the proton concentration was increased (< pH 6.0), and that such proton-evoked TRPV1 activation was observed in excised-membrane patches, indicating that protons gate TRPV1 directly (Tominaga et al., 1998). Thus, TRPV1 can be activated by at least three different pain-producing stimuli: capsaicin, heat (>43°C), or protons. These stimuli are likely to work in concert to regulate the activity of TRPV1 in vivo, especially under pathological conditions where tissue acidosis and elevated temperature may come into play. The studies outlined above provided in vitro evidence that TRPV1 serves as a transducer of noxious thermal and chemical stimuli in nociceptors. To determine whether TRPV1 really contributes to the detection of these noxious stimuli in vivo, mice lacking this protein were generated and analyzed for nociceptive function. Sensory neurons from mice lacking TRPV1 were deﬁcient in their responses to each

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of the reported noxious stimuli: capsaicin, protons, and heat (Caterina et al., 2000; Davis et al., 2000). Consistent with this observation, behavioral responses to capsaicin were absent and responses to acute thermal stimuli were diminished in these mice. The most prominent feature of the knockout mouse thermosensory phenotype was a virtual absence of thermal hypersensitivity in the setting of inﬂammation. These ﬁndings indicate that TRPV1 is essential for selective modalities of pain sensation and for tissue injury–induced thermal hyperalgesia. In addition, fever production in response to the bacterial pyrogen, lipopolysaccharide, was signiﬁcantly attenuated in TRPV1-deﬁcient mice, although its molecular mechanism is not known (Iida et al., 2005). The extent to which TRPV1 underlies the responses to noxious thermal stimuli and the contribution of other heat-sensitive channels remains to be clariﬁed. There was a drastic reduction of heat sensitivity in DRG neurons cultured from TRPV1-deﬁcient mice, but a small yet signiﬁcant percentage of DRG neurons showed large heat-evoked current responses to heat stimuli over 55°C. Furthermore, the TRPV1-deﬁcient mice showed impaired responses to noxious thermal stimuli only over 50°C, and a small but signiﬁcant amount of heat-evoked c-Fos induction persisted in spinal cord laminae I and II of TRPV1-deﬁcient mice (Caterina et al., 2000). These data supported the idea that other heat-sensitive channels contribute to the transmission and perception of high-intensity noxious thermal stimuli. Inﬂammatory pain is initiated by tissue damage/inﬂammation and is characterized by hypersensitivity both at the site of damage and in adjacent tissue. Stimuli that normally would not produce pain do so (allodynia), while previously noxious stimuli evoke even greater pain responses (hyperalgesia). One mechanism underlying these phenomena is the modulation (sensitization) of ion channels such as TRPV1 (Mizumura and Kumazawa, 1996; Wood and Perl, 1999; Woolf and Salter, 2000). Sensitization is triggered by extracellular inﬂammatory mediators that are released in vivo from surrounding damaged or inﬂamed tissues and from nociceptive neurons themselves (i.e., neurogenic inﬂammation). Mediators known to cause sensitization include prostaglandins, adenosine, serotonin, bradykinin, and ATP (Julius and Basbaum, 2001). Among the inﬂammatory mediators, extracellular ATP, bradykinin, prostagladins (prostaglandin E2 and prostaglandin I2), and trypsin or tryptase have been reported to potentiate TRPV1 responses through metabotropic P2Y2, B1 or B2, EP1 or IP, and proteinase-activated receptor-2 (PAR-2) receptors, respectively, in a PKC-dependent manner in both a heterologous expression system and native DRG neurons (Tominaga et al., 2001; Sugiura et al., 2002; Moriyama et al., 2003; Amadesi et al., 2004; Dai et al., 2004; Moriyama et al., 2005). In addition to potentiating capsaicin- or proton-evoked currents, ATP, bradykinin, prostaglandin E2, prostaglandin I2, trypsin, or tryptase also lower the temperature threshold for heat activation of TRPV1 to as low as 30°C, such that normally nonpainful thermal stimuli (i.e., normal body temperature) can activate TRPV1. Under these circumstances, those inﬂammatory mediators thus resemble direct activators of TRPV1. These inﬂammatory mediator-induced TRPV1-mediated hypersensitivities were conﬁrmed at the whole animal level using TRPV1-deﬁcient mice or mice lacking receptors for inﬂammatory mediators (Moriyama et al., 2003; Dai et al., 2004; Moriyama et al., 2005). PKC-dependent phosphorylation of TRPV1 has been found to be involved in the sensitization of TRPV1 by the inﬂammatory mediators. Indeed, two serine residues

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in the cytoplasmic domain of TRPV1 were identiﬁed as substrates for PKCdependent phosphorylation (Numazaki et al., 2002; Bhave et al., 2003). Although other pathways including a PKA-dependent one were found to be involved in sensitizing TRPV1, the PKC-dependent pathway seems to be predominantly involved in reducing the temperature threshold for TRPV1 activation. Tissue acidiﬁcation is induced in pathological conditions such as ischemia or inﬂammation (Bevan and Geppetti, 1994), and such acidiﬁcation exacerbates or causes pain. In addition to the direct activation of TRPV1, acidiﬁcation also shifts the temperatureresponse curve of TRPV1 to the left so that the channel can be activated at lower temperatures (lower than body temperature), and responses to heat are bigger at a given suprathreshold temperature (Tominaga et al., 1998). This phenomenon might also contribute to inﬂammatory pain. How can heat open this channel? A Q10 value of ∼26 for TRPV1 far surpasses the temperature dependence of the gating processes characterized by other ion channels (Q10 ∼3). The distal half of the TRPV1 carboxyl terminus was reported to be partially involved in thermal sensitivity (Vlachova et al., 2003). TRPV1 is known to have a voltage-dependent gating property (Gunthorpe et al., 2000). Nilius and colleagues have reported that temperature sensing in TRPV1 and TRPM8 (described later) is tightly linked to voltage-dependent gating (Voets et al., 2004). TRPV1 is activated upon depolarization, and changes in temperature result in graded shifts in its voltage-dependent activation curve. Through mathematical modeling, Nilius et al. made a single thermodynamic model that shows how temperature changes the thermal responsiveness of both channels. This suggests that amino acids responsible for voltage dependence are also involved in thermosensing, although the fourth TM domain of TRPV1 lacks the multiple positively charged residues typical of voltagegated channels.

TRPV2 (VRL-1) A protein with 49 percent of the same qualities as TRPV1 was isolated and designated vanilloid-receptor-like protein 1 (VRL-1) and later renamed TRPV2 (Caterina et al., 1999). TRPV2 is not activated by vanilloids, protons, or moderate thermal stimuli but can be activated by high temperatures with a threshold of ∼52°C. TRPV2 currents showed similar properties to those of TRPV1, such as an outwardly rectifying I-V relationship at positive potentials, inhibition by ruthenium red, and relatively high Ca2+ permeability (PCa/PNa= 2.9). However, whether TRPV2 is gated directly by high heat at a single-channel level is not known, probably because thermal noise inevitably contaminates recordings at such high temperatures. Intense TRPV2 immunoreactivity was observed in medium- to large-diameter cells in rat DRG neurons (Caterina et al., 1999; Ma, 2001; Ahluwalia et al., 2002; Lewinter et al., 2004). Many of the TRPV2 immunoreactive cells in rat DRG were co-stained with the anti-neuroﬁlament antibody N52, a marker for myelinated neurons. Dense TRPV2 immunoreactivity in the spinal cord was found in lamina I, inner lamina II, and laminae III/IV. This is consistent with the expression of TRPV2 in myelinated nociceptors that target laminae I and inner lamina II and in nonnociceptive Aβ ﬁbers that target laminae III/IV. Aδ mechano- and heat-sensitive

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(AMH) neurons in monkeys are medium- to large-diameter, lightly myelinated neurons that fall into two groups: type I AMHs have a heat threshold of ∼53°C, and type II AMHs are activated at 43°C (Treede et al., 1995). The TRPV2 localization data and the residual high temperature–evoked responses observed in the TRPV1deﬁcient mice suggest that TRPV2 expression might account for the high thermal threshold ascribed to type I AMH nociceptors. Temperatures activating TRPV2 are more harmful to our body than those activating TRPV1. Therefore, TRPV2 expression in the myelinated sensory ﬁbers seems reasonable because Aδ ﬁbers can transmit nociceptive information much faster than C ﬁbers, which exclusively express TRPV1. In addition to the acute thermal nociception, TRPV2 was suggested to be involved in peripheral sensitization during inﬂammation based on the result that injection of complete Freund’s adjuvant induced upregulation of TRPV2 expression in rat DRG neurons (Shimosato et al., 2005). The function of TRPV2 in Aβ ﬁbers is not known. TRPV2 transcript and protein were found not only in sensory neurons but also in motoneurons and in many nonneuronal tissues that are unlikely to be exposed to temperatures above 50°C (Caterina et al., 1999; Lewinter et al., 2004). These results indicate that TRPV2 undoubtedly contributes to numerous functions in addition to nociceptive processing. Indeed, growth factor-regulated channel, GRC (the likely mouse TRPV2 orthologue), was reported to translocate from cytosol to the plasma membrane upon stimulation with insulinlike growth factor (Kanzaki et al., 1999). And, mouse GRC has been found to be activated by stretching in cardiac myocytes (Iwata et al., 2003), suggesting that TRPV2 may be activated by mechanical stimuli.

WARM RECEPTORS TRPV3

AND

TRPV4

TRPV3 and TRPV4 (also known as VROAC or OTRPC4 from vanilloid receptor– related osmotically activated channel or OSM-9-like TRP channel 4, respectively) have been found to be activated by warm temperatures (∼34–38°C for TRPV3 and ∼27–35°C for TRPV4) in heterologous expression systems, and to be expressed in multiple tissues including, among others, sensory and hypothalamic neurons and keratinocytes (Guler et al., 2002; Peier, Reeve, et al., 2002; Smith et al., 2002; Watanabe et al., 2002; Xu et al., 2002). TRPV4 can alternatively be activated by nonthermal stimuli, including hypoosmolarity (Liedtke et al., 2000; Strotmann et al., 2000; Wissenbach et al., 2000), certain synthetic phorbol esters, and 5’,6’-epoxyeicosatrienoic acid (Watanabe et al., 2002; Watanabe, Vriens, et al., 2003). However, TRPV4 activation by heat was observed in a whole-cell conﬁguration but not in excised patches, suggesting the involvement of some cytosolic molecules in its activation, a different mechanism from that of TRPV1. Several approaches, including the knockdown of TRPV4 with gene disruption or antisense oligonucleotides, have led to reports that this protein is involved in mechanical stimulus– and hypotonicityinduced nociception in rodents at baseline or following hypersensitivity induced by

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prostaglandin injection or taxol neurotoxicity (Alessandri-Haber et al., 2003; Suzuki et al., 2003; Alessandri-Haber et al., 2004). In terms of the thermosensing ability of TRPV4 in vivo, peripheral nerve recordings suggested that there might be a decrease in warmth-evoked electrical activity in TRPV4-deﬁcient mice (Todaka et al., 2004). Moreover, both TRPV3 and TRPV4 have been reported to function in thermosenation by keratinocytes, based on studies of wild-type and TRPV4-deﬁcient keratinocytes (Chung et al., 2003; Chung et al., 2004). In contrast, no changes in escape latency from heat stimuli were observed in either the hot plate (Suzuki et al., 2003; Todaka et al., 2004) or radiant paw heating (Liedtke and Friedman, 2003) assays. After subcutaneous injection of capsaicin or carrageenan, however, TRPV4-deﬁcient mice showed longer escape latencies from a hot surface, relative to wild-type controls (Todaka et al., 2004). Thus, the precise contributions of TRPV4 to thermosensation and thermoregulation in vivo seem unclear. However, experiments using a thermal gradient apparatus revealed that TRPV4-deﬁcient mice selected warmer ﬂoor temperatures than wild-type mice (Lee et al., 2005). In addition, whereas wild-type mice failed to discriminate between ﬂoor temperatures of 30 and 34°C, TRPV4-deﬁcient mice exhibited a strong preference for 34°C and prolonged withdrawal latencies during acute tail heating. These results indicate that TRPV4 is required for normal thermal responsiveness. Further, the fact that TRPV4 is expressed in the hypothalamus, a center for body temperature regulation, suggests that TRPV4 is involved in thermoregulation in the brain by detecting temperature directly as well. TRPV3-deﬁcient mice lost preference for 35°C compared with room temperature, which was evident in wild-type mice, suggesting an important role for TRPV3 in innocuous thermosensation (Moqrich et al., 2005). In addition, TRPV3-deﬁcient mice exhibited delayed responses in tail ﬂick assay (over 50°C) and hot plate tests (over 55°C), indicating that TRPV3 is involved in thermal nociception, consistent with the sensitization of TRPV3 upon repeated noxious heat stimuli (Peier, Reeve, et al., 2002). The latter thermal nociceptive phenotype in TRPV3-deﬁcient mice is similar to those reported for TRPV1-deﬁcient mice, suggesting that these two TRPV channels have overlapping functions in vivo. The former innocuous thermal phenotype apparently seems to contradict one observed for TRPV4-deﬁcient mice. Functional discrimination of the two thermosensitive TRP channels with similar temperature activation thresholds in keratinocytes and the possible signal transduction from keratinocytes to sensory neurons would be important research directions in the future. Inﬂammatory mediator-induced thermal hyperalgesia was reported to be indistinguishable between wild-type and TRPV3-deﬁcient mice (Moqrich et al., 2005).

TRPM4

AND

TRPM5

TRPM4 is expressed in many tissues and cell types, and it functions as a Ca2+activated nonselective cation channel (Launay et al., 2002). Full-length TRPM4 (designated TRPM4b to distinguish it from the shorter TRPM4a variant) carries monovalent cations. Single-channel I-V relationship of TRPM4 is linear with a slope

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conductance of ∼25 pS, although voltage-dependent modulation causes an outwardly rectifying steady-state I-V relationship (Nilius et al., 2003). The gene encoding TRPM5 was identiﬁed during functional analysis of a chromosomal region that is associated with several tumors (Prawitt et al., 2000). TRPM5 is activated directly by elevated Ca2+ (Hofmann et al., 2003; Prawitt et al., 2003), and it was reported to be activated in the downstream of activation of G-protein-coupled taste receptors (Perez et al., 2002). Like TRPM4, TRPM5 is a monovalent-speciﬁc ion channel with ∼25 pS conductance and has the steady-state I-V relationship characterized by strong outward rectiﬁcation. TRPM4 and TRPM5 have been found to be temperature-sensitive, heat-activated ion channels (Talavera et al., 2005). TRPM4- or TRPM5-mediated inward currents increased steeply at temperatures between 15 and 35°C. Heat activation was due to a temperature-dependent shift of the activation curve, in analogy to TRPV1. Also, increasing temperature between 15 and 35°C markedly enhanced the gustatory nerve response to sweet compounds in wild-type mice but not in mice lacking TRPM5, suggesting that temperature sensitivity of TRPM5 may underlie known effects of temperature on perceived taste in humans (Bartoshuk et al., 1982; Green and Frankmann, 1988).

COLD RECEPTORS TRPM8 (CMR1) A distinct class of cold-sensitive ﬁbers has been described as polymodal nociceptors, responding to noxious cold, heat, and pinching (Campero et al., 1996; Simone and Kajander, 1997; Cain et al., 2001), although an early study suggested a distinction between cold sensation and painful sensation (Klement and Arndt, 1992). The cooling sensation of menthol, a chemical agent found in mint, is well established (Hensel and Zoterman, 1951), and both cooling and menthol have been suggested to be transduced through a nonselective cation channel in DRG neurons (Reid and Flonta, 2001; Reid et al., 2002). Two groups independently cloned and characterized a cold receptor, TRPM8 (also known as CMR1, for cold- and menthol-sensitive receptor 1), which can also be activated by menthol (McKemy et al., 2002; Peier, Moqrich, et al., 2002). In heterologous expression systems, TRPM8 could be activated by menthol or by cooling, with an activation temperature of ∼25–28°C. TRPM8 could alternatively be activated by other cooling compounds, such as menthone, eucalyptol, and icilin. There also appears to be interaction between effective stimuli for TRPM8, in that subthreshold concentrations of menthol increased the temperature threshold for TRPM8 activation from 25°C to 30°C. This is reminiscent of TRPV1, whose activation temperature is reduced under mildly acidic conditions that do not open TRPV1 alone (Tominaga et al., 1998). Whole-cell recording in HEK293 cells expressing TRPM8 revealed that TRPM8 is a nonselective cation channel with relatively high Ca2+ permeability (PCa/PNa= 3.3) and that TRPM8 shows an outwardly rectifying I-V relationship like TRPV1. Single-channel recordings showed a conductance of 83 pS at positive potentials. Cold activation of TRPM8 was reported to

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be due to a temperature-dependent shift of the activation curve, in analogy to TRPV1, TRPM4, and TRPM5 (Voets et al., 2004). TRPM8 is expressed in a subset of DRG and TG neurons that can be classiﬁed as small-diameter C ﬁbers (McKemy et al., 2002; Peier, Moqrich, et al., 2002; Nealen et al., 2003). Interestingly, however, TRPM8 is not coexpressed with TRPV1, which marks a class of nociceptors. Whether TRPM8 is involved in cold nociception remains to be clariﬁed, although inﬂammatory mediators were found to downregulate TRPM8 via PKC-mediated phosphorylation (Premkumar et al., 2005), suggesting that the TRPM8 downregulation might aggravate thermal hyperalgesia in the context of inﬂammation. Regarding channel regulation, reciprocal modulation of TRPM8 and TRPV1 by PIP2 has been described (Chuang et al., 2001; Liu and Qin, 2005; Rohacs et al., 2005). TRPM8 is activated, whereas TRPV1 is tonically inhibited by PIP2. Hydrolysis of PIP2 by activating phospholipase C downregulates TRPM8 function and upregulates TRPV1 function. Bradykinin-induced downregulation of TRPM8 and upregulation of TRPV1 could almost completely be reversed by a PKC inhibitor (Premkumar et al., 2005). Thus, this mechanism of TRPM8 downregulation could worsen inﬂammatory hyperalgesia because more cooling would be necessary to activate TRPM8.

TRPA1 (ANKTM1) TRPA1 (also known as ANKTM1, a channel containing both N-terminal ankyrin repeat domains and six transmembrane domains) was reported as a distantly related TRP channel that is activated by cold with a lower activation threshold as compared to TRPM8 (Story et al., 2003). In heterologous expression systems, TRPA1 was activated by cold stimuli with an activation temperature of about 17°C, which is close to the reported noxious cold threshold. This led to the suggestion that TRPA1 is involved in cold nociception. Indeed, TRPA1 has been involved in cold hyperalgesia caused by inﬂammation or nerve injury via activation of MAP kinase (Obata et al., 2005). Whole-cell recording in ﬁbroblastic CHO cells expressing TRPA1 revealed cationic permeability with similar preferences for monovalent and divalent cations (PCa/PNa= 0.8) and an outwardly rectifying I-V relationship. Whether TRPA1 is gated directly by cold remains to be elucidated because no single-channel data have been reported. A recent study from another group failed to reproduce cold responsiveness in TRPA1 (Jordt et al., 2004). The reason for this apparent discrepancy is unclear. However, both groups have demonstrated that TRPA1 can be activated by pungent isothiocyanate compounds such as those found in wasabi, horseradish, and mustard oil (Bandell et al., 2004; Jordt et al., 2004) or by allicin, a pungent ingredient of garlic (Bautista et al., 2005; Macpherson et al., 2005). Thus, several of the thermosensitive TRP channels likely to be involved in nociception can be activated by stimuli other than temperature (Table 20.1). Unlike TRPM8, TRPA1 is speciﬁcally expressed in a subset of sensory neurons that express the nociceptive markers CGRP and substance P (Story et al., 2003). Furthermore, TRPA1 is frequently coexpressed with TRPV1, raising the possibility that TRPA1 and TRPV1 mediate the function of a class of polymodal nociceptors.

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TABLE 20.1 The Eight Thermosensitive TRP Channels

Receptor

Temperature Threshold for Activation

Expression

Other Effective Stimuli

TRPV1

43°C <

Sensory neurons Epithelial cells

Capsaicin/proton Lipids/allicin 2-aminoethoxydiphenyl borate

TRPV2

52°C <

Sensory neurons/brain/spinal cord Lung/liver/spleen/colon/heart

Mechanical stimulus 2-aminoethoxydiphenyl borate

TRPV3

32–39°C <

Sensory neurons/brain Spinal cord/skin/stomach/colon

Camphor 2-aminoethoxydiphenyl borate

TRPV4

27–35°C <

Sensory neurons/hypothalamus Skin/kidney/lung/inner ear

Hypotonic stimulus/4PDD Mechanical stimulus Epoxyeicosatrienoic acids

TRPM4

15~35°C

Ubiquitous

TRPM5

15~35°C

Taste cells/pancreas

TRPM8

< 25–28°C

Sensory neurons

TRPA1

< 17°C

Sensory neurons/inner ear

Ca2+ Ca2+/Phospholipase C Menthol Allyl isothiocyanate Δ9-tetrahydrocannabinol Cinnamaldehyde/allicin Mechanical stimulus

Such coexpression might also explain the paradoxical hot sensation experienced when one is exposed to a very cold stimulus.

THERMAL NOCICEPTION THROUGH TRP CHANNELS IN INVERTEBRATES Invertebrates, like vertebrates, need to detect environmental temperature for their survival and to escape potentially dangerous stimuli. Interestingly, recent experiments indicate that TRP channels play a critical role in thermosensation in invertebrate, as well as vertebrate, species (Goodman, 2003; Zars, 2003). A TRPA1 homologue identiﬁed in Drosophila, named Painless, is expressed in thermosensory neurons and was found to be required for withdrawal responses to noxious thermal and mechanical stimuli. Although it is not clear whether Painless senses temperature directly (Tracey et al., 2003), it may be noteworthy that the threshold for evoking Painless-dependent behavior in Drosophila larvae is approximately 39–42°C, similar to the threshold for thermal pain in mammals. Moreover, residual withdrawal responses to a 52°C stimulus are preserved in painless mutants, suggesting that Drosophila, like mammals, may possess multiple mechanisms for thermonociception. Another recent study showed that the apparent Drosophila ortholog of TRPA1

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(not Painless) is activated by warm temperatures (about 24–29°C) when expressed heterologously (Viswanath et al., 2003). Given that these species prefer an ambient temperature of ~24°C, this warmth-sensitive TRPA1 orthologue might also participate in some form of aversive heat sensation. It is particularly intriguing that the mammalian and Drosophila TRPA1 orthologues, while similar in primary sequence, are activated by two apparently opposite stimuli: cold and warmth. The mechanistic basis of this difference remains to be determined.

CONCLUSION Signiﬁcant advances in thermosensation and thermoregulation research have been made in the last several years with the cloning and characterization of thermosensitive TRP channels. With these clones, we can understand thermosensation and thermoregulation from a molecular standpoint. How temperature gates TRP channels is a very interesting question, but is not answered yet. Chemical agonists have been identiﬁed for many of the thermosensitive TRP channels (TRPV1, TRPV2, TRPV3, TRPV4, TRPM8, and TRPA1) (Table 20.1). Understanding the mechanisms of direct or indirect TRP channel opening by chemical activators might clarify the gating mechanism by thermal stimulation. It is also not known how multiple thermosensitive TRP channels are used within the same or different sensory neurons to provide us with the ability to evaluate temperature precisely over a broad range. Central integration of the thermal information acquired by peripheral nerve endings is almost certainly a critical component of this process. In addition, function of thermosensitive TRP channels in tissues other than neurons will have to be extensively investigated. Furthermore, regulatory mechanisms similar to those emerging for TRPV1 are likely to exist for most of the thermosensitive TRP channels. Therefore, an understanding of the molecules surrounding thermosensitive TRP channels will be indispensable to a complete understanding of thermosensation and thermoregulation.

REFERENCES Ahluwalia J, Rang H, and Nagy I. 2002. The putative role of vanilloid receptor-like protein-1 in mediating high-threshold noxious heat-sensitivity in rat cultured primary sensory neurons. Eur J Neurosci 16:1483–1489. Alessandri-Haber N, Dina OA, Yeh JJ, Parada CA, Reichling DB, and Levine JD. 2004. Transient receptor potential vanilloid 4 is essential in chemotherapy-induced neuropathic pain in the rat. J Neurosci 24:4444–4452. Alessandri-Haber N, Yeh JJ, Boyd AE, Parada CA, Chen X, Reichling DB, and Levine JD. 2003. Hypotonicity induces TRPV4-mediated nociception in rat. Neuron 39: 497–511. Amadesi S, Nie J, Vergnolle N, Cottrell GS, Grady EF, Trevisani M, Manni C, Geppetti P, McRoberts JA, Ennes H, Davis JB, Mayer EA, and Bunnett NW. 2004. Proteaseactivated receptor 2 sensitizes the capsaicin receptor transient receptor potential vanilloid receptor 1 to induce hyperalgesia. J Neurosci 24:4300–4312. Bandell M, Story GM, Hwang SW, Viswanath V, Eid SR, Petrus MJ, Earley TJ, and Patapoutian A. 2004. Noxious cold ion channel TRPA1 is activated by pungent compounds and bradykinin. Neuron 41:849–857.

282

TRP Ion Channel Function in Sensory Transduction

Bartoshuk LM, Rennert K, Rodin J, and Stevens JC. 1982. Effects of temperature on the perceived sweetness of sucrose. Physiol Behav 28:905–910. Bautista DM, Movahed P, Hinman A, Axelsson HE, Sterner O, Hogestatt ED, Julius D, Jordt SE, and Zygmunt PM. 2005. Pungent products from garlic activate the sensory ion channel TRPA1. Proc Natl Acad Sci USA 102:12248–12252. Bevan S and Geppetti P. 1994. Protons: small stimulants of capsaicin-sensitive sensory nerves. Trends Neurosci 17:509–512. Bhave G, Hu HJ, Glauner KS, Zhu W, Wang H, Brasier DJ, Oxford GS, and Gereau RWt. 2003. Protein kinase C phosphorylation sensitizes but does not activate the capsaicin receptor transient receptor potential vanilloid 1 (TRPV1). Proc Natl Acad Sci USA 100:12480–12485. Cain DM, Khasabov SG, and Simone DA. 2001. Response properties of mechanoreceptors and nociceptors in mouse glabrous skin: an in vivo study. J Neurophysiol 85:1561– 1574. Campero M, Serra J, and Ochoa JL. 1996. C-polymodal nociceptors activated by noxious low temperature in human skin. J Physiol 497 (Pt. 2):565–572. Caterina MJ and Julius D. 2001. The vanilloid receptor: a molecular gateway to the pain pathway. Annu Rev Neurosci 24:487–517. Caterina MJ, Lefﬂer A, Malmberg AB, Martin WJ, Trafton J, Petersen-Zeitz KR, Koltzenburg M, Basbaum AI, and Julius D. 2000. Impaired nociception and pain sensation in mice lacking the capsaicin receptor. Science 288:306–313. Caterina MJ, Rosen TA, Tominaga M, Brake AJ, and Julius D. 1999. A capsaicin-receptor homologue with a high threshold for noxious heat. Nature 398:436–441. Caterina MJ, Schumacher MA, Tominaga M, Rosen TA, Levine JD, and Julius D. 1997. The capsaicin receptor: a heat-activated ion channel in the pain pathway. Nature 389:816–824. Cesare P, Dekker LV, Sardini A, Parker PJ, and McNaughton PA. 1999. Speciﬁc involvement of PKC-epsilon in sensitization of the neuronal response to painful heat. Neuron 23:617–624. Cesare P and McNaughton P. 1996. A novel heat-activated current in nociceptive neurons and its sensitization by bradykinin. Proc Natl Acad Sci USA 93:15435–15439. Chuang HH, Prescott ED, Kong H, Shields S, Jordt SE, Basbaum AI, Chao MV, and Julius D. 2001. Bradykinin and nerve growth factor release the capsaicin receptor from PtdIns(4,5)P2-mediated inhibition. Nature 411:957–962. Chung MK, Lee H, and Caterina MJ. 2003. Warm temperatures activate TRPV4 in mouse 308 keratinocytes. J Biol Chem 278:32037–32046. Chung MK, Lee H, Mizuno A, Suzuki M, and Caterina MJ. 2004. TRPV3 and TRPV4 mediate warmth-evoked currents in primary mouse keratinocytes. J Biol Chem 279:21569– 21575. Dai Y, Moriyama T, Higashi T, Togashi K, Kobayashi K, Yamanaka H, Tominaga M, and Noguchi K. 2004. Proteinase-activated receptor 2-mediated potentiation of transient receptor potential vanilloid subfamily 1 activity reveals a mechanism for proteinaseinduced inﬂammatory pain. J Neurosci 24:4293–4299. Davis JB, Gray J, Gunthorpe MJ, Hatcher JP, Davey PT, Overend P, Harries MH, Latcham J, Clapham C, Atkinson K, Hughes SA, Rance K, Grau E, Harper AJ, Pugh PL, Rogers DC, Bingham S, Randall A, and Sheardown SA. 2000. Vanilloid receptor-1 is essential for inﬂammatory thermal hyperalgesia. Nature 405:183–187. Goodman MB. 2003. Sensation is painless. Trends Neurosci 26:643–645. Green BG and Frankmann SP. 1988. The effect of cooling on the perception of carbohydrate and intensive sweeteners. Physiol Behav 43:515–519.

The Role of TRP Channels in Thermosensation

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Guler AD, Lee H, Iida T, Shimizu I, Tominaga M and Caterina M. 2002. Heat-evoked activation of the ion channel, TRPV4. J Neurosci 22:6408–6414. Gunthorpe MJ, Harries MH, Prinjha RK, Davis JB, and Randall A. 2000. Voltage- and timedependent properties of the recombinant rat vanilloid receptor (rVR1). J Physiol 525 (Pt. 3):747–759. Hensel H and Zoterman Y. 1951. The effect of menthol on the thermoreceptors. Acta Physiol Scand 24:27–34. Hofmann T, Chubanov V, Gudermann T, and Montell C. 2003. TRPM5 is a voltage-modulated and Ca(2+)-activated monovalent selective cation channel. Curr Biol 13:1153– 1158. Iida T, Shimizu I, Nealen ML, Campbell A, and Caterina M. 2005. Attenuated fever response in mice lacking TRPV1. Neurosci Lett 378:28–33. Iwata Y, Katanosaka Y, Arai Y, Komamura K, Miyatake K, and Shigekawa M. 2003. A novel mechanism of myocyte degeneration involving the Ca2+-permeable growth factor– regulated channel. J Cell Biol 161:957–967. Jordt SE, Bautista DM, Chuang HH, McKemy DD, Zygmunt PM, Hogestatt ED, Meng ID, and Julius D. 2004. Mustard oils and cannabinoids excite sensory nerve ﬁbres through the TRP channel ANKTM1. Nature 427:260–265. Jordt SE, McKemy DD, and Julius D. 2003. Lessons from peppers and peppermint: the molecular logic of thermosensation. Curr Opin Neurobiol 13:487–492. Julius D and Basbaum AI. 2001. Molecular mechanisms of nociception. Nature 413:203–210. Kanzaki M, Zhang YQ, Mashima H, Li L, Shibata H, and Kojima I. 1999. Translocation of a calcium-permeable cation channel induced by insulin-like growth factor-I. Nat Cell Biol 1:165–170. Kirschstein T, Busselberg D, and Treede RD. 1997. Coexpression of heat-evoked and capsaicin-evoked inward currents in acutely dissociated rat dorsal root ganglion neurons. Neurosci Lett 231:33–36. Klement W and Arndt JO. 1992. The role of nociceptors of cutaneous veins in the mediation of cold pain in man. J Physiol 449:73–83. LaMotte RH and Campbell JN. 1978. Comparison of responses of warm and nociceptive Cﬁber afferents in monkeys with human judgments of thermal pain. J Neurophysiol 41:509–528. Launay P, Fleig A, Perraud AL, Scharenberg AM, Penner R, and Kinet JP. 2002. TRPM4 is a Ca2+-activated nonselective cation channel mediating cell membrane depolarization. Cell 109:397–407. Lee H, Iida T, Mizuno A, Suzuki M, and Caterina MJ. 2005. Altered thermal selection behavior in mice lacking transient receptor potential vanilloid 4. J Neurosci 25:1304– 1310. Lewinter RD, Skinner K, Julius D, and Basbaum AI. 2004. Immunoreactive TRPV-2 (VRL-1), a capsaicin receptor homolog, in the spinal cord of the rat. J Comp Neurol 470:400–408. Liedtke W, Choe Y, Marti-Renom MA, Bell AM, Denis CS, Sali A, Hudspeth AJ, Friedman JM, and Heller S. 2000. Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor. Cell 103:525–535. Liedtke W and Friedman JM. 2003. Abnormal osmotic regulation in trpv4-/- mice. Proc Natl Acad Sci USA 100:13698–13703. Liu B and Qin F. 2005. Functional control of cold- and menthol-sensitive TRPM8 ion channels by phosphatidylinositol 4,5-bisphosphate. J Neurosci 25:1674–1681. Ma QP. 2001. Vanilloid receptor homologue, VRL1, is expressed by both A- and C-ﬁber sensory neurons. Neuroreport 12:3693–3695.

284

TRP Ion Channel Function in Sensory Transduction

Macpherson LJ, Geierstanger BH, Viswanath V, Bandell M, Eid SR, Hwang S, and Patapoutian A. 2005. The pungency of garlic: activation of TRPA1 and TRPV1 in response to allicin. Curr Biol 15:929–934. McKemy DD, Neuhausser WM, and Julius D. 2002. Identiﬁcation of a cold receptor reveals a general role for TRP channels in thermosensation. Nature 416:52–58. Mizumura K and Kumazawa T. 1996. Modiﬁcation of nociceptor responses by inﬂammatory mediators and second messengers implicated in their action—a study in canine testicular polymodal receptors. Prog Brain Res 113:115–141. Moqrich A, Hwang SW, Earley TJ, Petrus MJ, Murray AN, Spencer KS, Andahazy M, Story GM, and Patapoutian A. 2005. Impaired thermosensation in mice lacking TRPV3, a heat and camphor sensor in the skin. Science 307:1468–1472. Moriyama T, Higashi T, Togashi K, Iida T, Segi E, Sugimoto Y, Tominaga T, Narumiya S, and Tominaga M. 2005. Sensitization of TRPV1 by EP1 and IP reveals peripheral nociceptive mechanism of prostaglandins. Molecular Pain 1:3–12. Moriyama T, Iida T, Kobayashi K, Higashi T, Fukuoka T, Tsumura H, Leon C, Suzuki N, Inoue K, Gachet C, Noguchi K, and Tominaga M. 2003. Possible involvement of P2Y2 metabotropic receptors in ATP-induced transient receptor potential vanilloid receptor 1-mediated thermal hypersensitivity. J Neurosci 23:6058–6062. Nagy I and Rang HP. 1999. Similarities and differences between the responses of rat sensory neurons to noxious heat and capsaicin. J Neurosci 19:10647–10655. Nealen ML, Gold MS, Thut PD, and Caterina MJ. 2003. TRPM8 mRNA is expressed in a subset of cold-responsive trigeminal neurons from rat. J Neurophysiol 90:515– 520. Nilius B, Prenen J, Droogmans G, Voets T, Vennekens R, Freichel M, Wissenbach U, and Flockerzi V. 2003. Voltage dependence of the Ca2+-activated cation channel TRPM4. J Biol Chem 278:30813–30820. Numazaki M, Tominaga T, Toyooka H, and Tominaga M. 2002. Direct phosphorylation of capsaicin receptor VR1 by protein kinase C epsilon and identiﬁcation of two target serine residues. J Biol Chem 277:13375–13378. Obata K, Katsura H, Mizushima T, Yamanaka H, Kobayashi K, Dai Y, Fukuoka T, Tokunaga A, Tominaga M, and Noguchi K. 2005. TRPA1 induced in sensory neurons contributes to cold hyperalgesia after inﬂammation and nerve injury. J Clin Invest 115:2393– 2401. Patapoutian A, Peier AM, Story GM, and Viswanath V. 2003. ThermoTRP channels and beyond: mechanisms of temperature sensation. Nat Rev Neurosci 4:529–539. Peier AM, Moqrich A, Hergarden AC, Reeve AJ, Andersson DA, Story GM, Earley TJ, Dragoni I, McIntyre P, Bevan S, and Patapoutian A. 2002. A TRP channel that senses cold stimuli and menthol. Cell 108:705–715. Peier AM, Reeve AJ, Andersson DA, Moqrich A, Earley TJ, Hergarden AC, Story GM, Colley S, Hogenesch JB, McIntyre P, Bevan S, and Patapoutian A. 2002. A heat-sensitive TRP channel expressed in keratinocytes. Science 296:2046–2049. Perez CA, Huang L, Rong M, Kozak JA, Preuss AK, Zhang H, Max M, and Margolskee RF. 2002. A transient receptor potential channel expressed in taste receptor cells. Nat Neurosci 5:1169–1176. Prawitt D, Enklaar T, Klemm G, Gartner B, Spangenberg C, Winterpacht A, Higgins M, Pelletier J, and Zabel B. 2000. Identiﬁcation and characterization of MTR1, a novel gene with homology to melastatin (MLSN1) and the trp gene family located in the BWS-WT2 critical region on chromosome 11p15.5 and showing allele-speciﬁc expression. Hum Mol Genet 9:203–216.

The Role of TRP Channels in Thermosensation

285

Prawitt D, Monteilh-Zoller MK, Brixel L, Spangenberg C, Zabel B, Fleig A, and 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. Premkumar LS, Raisinghani M, Pingle SC, Long C, and Pimentel F. 2005. Downregulation of transient receptor potential melastatin 8 by protein kinase C–mediated dephosphorylation. J Neurosci 25:11322–11329. Reichling DB and Levine JD. 1997. Heat transduction in rat sensory neurons by calciumdependent activation of a cation channel. Proc Natl Acad Sci USA 94:7006–7011. Reid G, Babes A, and Pluteanu F. 2002. A cold- and menthol-activated current in rat dorsal root ganglion neurones: properties and role in cold transduction. J Physiol 545:595– 614. Reid G and Flonta ML. 2001. Physiology. Cold current in thermoreceptive neurons. Nature 413:480. Rohacs T, Lopes CM, Michailidis I, and Logothetis DE. 2005. PI(4,5)P2 regulates the activation and desensitization of TRPM8 channels through the TRP domain. Nat Neurosci 8:626–634. Shimosato G, Amaya F, Ueda M, Tanaka Y, Decoster I, and Tanaka M. 2005. Peripheral inﬂammation induces up-regulation of TRPV2 expression in rat DRG. Pain 119:225– 232. Simone DA and Kajander KC. 1997. Responses of cutaneous A-ﬁber nociceptors to noxious cold. J Neurophysiol 77:2049–2060. Smith GD, Gunthorpe MJ, Kelsell RE, Hayes PD, Reilly P, Facer P, Wright JE, Jerman JC, Walhin JP, Ooi L, Egerton J, Charles KJ, Smart D, Randall AD, Anand P, and Davis JB. 2002. TRPV3 is a temperature-sensitive vanilloid receptor-like protein. Nature 418:186–190. Story GM, Peier AM, Reeve AJ, Eid SR, Mosbacher J, Hricik TR, Earley TJ, Hergarden AC, Andersson DA, Hwang SW, McIntyre P, Jegla T, Bevan S, and Patapoutian A. 2003. ANKTM1, a TRP-like channel expressed in nociceptive neurons, is activated by cold temperatures. Cell 112:819–829. Strotmann R, Harteneck C, Nunnenmacher K, Schultz G, and Plant TD. 2000. OTRPC4, a nonselective cation channel that confers sensitivity to extracellular osmolarity. Nat Cell Biol 2:695–702. Sugiura T, Tominaga M, Katsuya H, and Mizumura K. 2002. Bradykinin lowers the threshold temperature for heat activation of vanilloid receptor 1. J Neurophysiol 88:544–548. Suzuki M, Mizuno A, Kodaira K, and Imai M. 2003. Impaired pressure sensation in mice lacking TRPV4. J Biol Chem 278:22664–22668. Talavera K, Yasumatsu K, Voets T, Droogmans G, Shigemura N, Ninomiya Y, Margolskee RF, and Nilius B. 2005. Heat activation of TRPM5 underlies thermal sensitivity of sweet taste. Nature 438:1022–1025. Tillman DB, Treede RD, Meyer RA, and Campbell JN. 1995. Response of C ﬁbre nociceptors in the anaesthetized monkey to heat stimuli: estimates of receptor depth and threshold. J Physiol 485 (Pt. 3):753–765. Todaka H, Taniguchi J, Satoh J, Mizuno A, and Suzuki M. 2004. Warm temperature-sensitive transient receptor potential vanilloid 4 (TRPV4) plays an essential role in thermal hyperalgesia. J Biol Chem 279:35133–35138. Tominaga M and Caterina MJ. 2004. Thermosensation and pain. J Neurobiol 61:3–12. Tominaga M, Caterina MJ, Malmberg AB, Rosen TA, Gilbert H, Skinner K, Raumann BE, Basbaum AI, and Julius D. 1998. The cloned capsaicin receptor integrates multiple pain-producing stimuli. Neuron 21:531–543.

286

TRP Ion Channel Function in Sensory Transduction

Tominaga M, Wada M, and Masu M. 2001. Potentiation of capsaicin receptor activity by metabotropic ATP receptors as a possible mechanism for ATP-evoked pain and hyperalgesia. Proc Natl Acad Sci USA 98:6951–6956. Tracey WD, Jr., Wilson RI, Laurent G, and Benzer S. 2003. painless, a Drosophila gene essential for nociception. Cell 113:261–273. Treede RD, Meyer RA, Raja SN, and Campbell JN. 1995. Evidence for two different heat transduction mechanisms in nociceptive primary afferents innervating monkey skin. J Physiol 483 (Pt. 3):747–758. Viswanath V, Story GM, Peier AM, Petrus MJ, Lee VM, Hwang SW, Patapoutian A, and Jegla T. 2003. Opposite thermosensor in fruitﬂy and mouse. Nature 423:822–823. Vlachova V, Teisinger J, Susankova K, Lyfenko A, Ettrich R, and Vyklicky L. 2003. Functional role of C-terminal cytoplasmic tail of rat vanilloid receptor 1. J Neurosci 23:1340– 1350. Voets T, Droogmans G, Wissenbach U, Janssens A, Flockerzi V, and Nilius B. 2004. The principle of temperature-dependent gating in cold- and heat-sensitive TRP channels. Nature 430:748–754. Watanabe H, Davis JB, Smart D, Jerman JC, Smith GD, Hayes P, Vriens J, Cairns W, Wissenbach U, Prenen J, Flockerzi V, Droogmans G, Benham CD, and Nilius B. 2002. Activation of TRPV4 channels (hVRL-2/mTRP12) by phorbol derivatives. J Biol Chem 277:13569–13577. Watanabe H, Vriens J, Prenen J, Droogmans G, Voets T, and Nilius B. 2003. Anandamide and arachidonic acid use epoxyeicosatrienoic acids to activate TRPV4 channels. Nature 424:434–438. Watanabe H, Vriens J, Suh SH, Benham CD, Droogmans G, and Nilius B. 2002. Heat-evoked activation of TRPV4 channels in a HEK293 cell expression system and in native mouse aorta endothelial cells. J Biol Chem 277:47044–47051. Wissenbach U, Bodding M, Freichel M, and Flockerzi V. 2000. Trp12, a novel Trp-related protein from kidney. FEBS Lett 485:127–134. Wood JN and Perl ER. 1999. Pain. Curr Opin Genet Dev 9:328–332. Woolf CJ and Salter MW. 2000. Neuronal plasticity: increasing the gain in pain. Science 288:1765–1769. Xu H, Ramsey IS, Kotecha SA, Moran MM, Chong JA, Lawson D, Ge P, Lilly J, SilosSantiago I, Xie Y, DiStefano PS, Curtis R, and Clapham DE. 2002. TRPV3 is a calcium-permeable temperature-sensitive cation channel. Nature 418:181–186. Zars T. 2003. Hot and cold in Drosophila larvae. Trends Neurosci 26:575–577.

21

Voltage and Temperature Gating of ThermoTRP Channels Ramon Latorre Guillermo Vargas Gerardo Orta Centro de Estudios Cientiﬁcos

Sebastian Brauchi Universidad Austral de Chile and Centro de Estudios Cientiﬁcos

CONTENTS Abstract ..................................................................................................................287 Introduction............................................................................................................288 Heat Receptors...........................................................................................288 Cold Receptors ..........................................................................................288 ThermoTRP Biophysics ........................................................................................289 Temperature Dependence ..........................................................................289 Voltage Dependence ..................................................................................291 Protein Denaturation, Lipids, and ThermoTRP Channels ....................................294 Multistate Nature of ThermoTRPs’ Opening Process ..........................................295 A Minimum Model That Explains the Change in Length of Bursts and Gaps with Temperature...................................................................296 An Allosteric Model for ThermoTRP Activation .................................................297 Testing the Allosteric Model .....................................................................297 References..............................................................................................................299

ABSTRACT In this chapter we address the puzzle of thermoTRP channel temperature and voltage sensitivity from a biophysical point of view. We deal with the problem through a classic thermodynamic approach, followed by a comparison of thermoTRPs’ behavior

287

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with our present knowledge regarding protein structure folding. On the basis of their temperature dependence and voltage dependence, we propose a general allosteric model for thermoTRP channel gating.

INTRODUCTION HEAT RECEPTORS TRP ion channels are one of the largest groups of ion channels, and this group is composed of six protein families.1–5 With few exceptions, TRP channel subunits are predicted to have six transmembrane domains, and a subset of these channels, the thermoTRPs, is activated by distinct physiological temperatures. ThermoTRP channels have a tetrameric structure in which the subunits preferentially assemble into homomeric complexes.6,7 Six members of the mammalian TRP ion channels respond to varied temperature thresholds. TRPV1, TRPV2, TRPV3, and TRPV4 are heat activated, whereas TRPM8 and TRPA1 are activated by cold.8–10 The ﬁrst member of this family to be cloned was the vanilloid receptor 1 (TRPV1),11 which is mainly expressed in small-diameter sensory neurons of the trigeminal (TG) and dorsal root ganglia (DRG). Capsaicin (the active component of “hot” chili peppers), pH, and noxious thermal stimuli (~43°C) activate TRPV1.12 TRPV1 can be activated at room temperature when the pH is decreased to <6. This reduction in the temperature threshold for activation could be the mechanism involved in inﬂammatory pain because tissue acidiﬁcation is induced in ischemia and inﬂammation.12 More recently, Macpherson et al.13 showed that allicin, a component of fresh garlic, is able to activate both TRPV1 and TRPA1 and is likely responsible for garlic’s pungency. Thus, TRPV1 can act as a transducer of noxious thermal and chemical stimuli in nociceptors, as has been conﬁrmed in mice lacking this protein.14 TRPV2, present in medium- to large-diameter DRG neurons and in motor neurons, is activated by high temperatures with a threshold of about 52°C.15–18 TRPV3 is expressed in sensory neurons and keratinocytes.19–21 It is activated by innocuous temperatures (~33°C), and its activity is further increased at noxious temperatures. TRPV3 is also markedly sensitized by repeated heat stimuli, but the mechanism accounting for this phenomenon is unknown;9 the gradually sensitized initial phase is followed by an abrupt increase in current.22 More recently, Moqrich et al.23 found that the natural compound camphor specifically activates TRPV3 channels. TRPV4, previously known to be activated by osmotic pressure, was also shown to be activated by warm temperature stimuli (temperature threshold of ~35°C) and is expressed in DRGs, neurons, and keratinocytes, among other tissues.24–29

COLD RECEPTORS TRPM8, which was cloned independently by two groups and characterized as a cold- and menthol-sensitive receptor, is expressed selectively in a small population of primary sensory neurons.30,31 It is activated directly by gentle cooling (threshold ∼22–30°C) and depolarizes sensory neurons.10,32 TRPM8 is activated by menthol,

Voltage and Temperature Gating of ThermoTRP Channels

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the super-cooling agent icilin, and eucalyptol, albeit with lower efﬁcacy and potency than menthol.30 Strong cooling activates (<17°C) TRPA1 channels and has been suggested to be important in the transduction of strong (painful) cooling stimuli.33 Isothiocyanate compounds, the active components of wasabi and mustard, activate TRPA1 channels, identifying this channel as the molecular target for the pungent action of mustard oils.34,35 However, the involvement of this channel in cold sensitivity of sensory neurons has been questioned.36,34 Bandell et al.35 reported that noxious cold activates TRPA1 channels both in heterologous system and DRG cultures. Thus, thermoTRPs provide us with the ability to evaluate temperature precisely over a wide range of temperatures, including those considered noxious. Their behavior suggests that either they are direct temperature sensors or they are activated by some ubiquitous temperature-sensitive molecular component extrinsic to the channel-forming protein. It is of importance therefore to unveil the molecular mechanisms that make these proteins so exquisitely temperature sensitive.

THERMOTRP BIOPHYSICS TEMPERATURE DEPENDENCE Channel gating is known to be affected by temperature. Q10 value is used to estimate the temperature dependence of a given system, taking the activity of the system at two different temperatures separated by 10°C. In the case of thermoTRPs, the Q10 could be easily obtained from the macroscopic currents using the following deﬁnition: ⎛I ⎞ Q10 = ⎜ 2 ⎟ ⎝ I1 ⎠

10

( T2 − T1 )

(1)

where I1 and I2 are the obtained currents at given temperatures T1 and T2. Although the Q10 obtained by this procedure is very useful as a comparative value, it lacks thermodynamic meaning. A relationship between Q10 and the Arrehenius activation energy (Ea) can be obtained by measuring rate constants, k, at different temperatures since: k = Ae

− Ea RT

(2)

where A is a constant called the frequency factor. In the case of a ﬁrst-order reaction k = 1/ where τ is the time constant it is easy to show that ⎛ TT ⎞ τ Ea = R ⎜ 1 2 ⎟ × ln 1 ⎜⎝ T − T ⎟⎠ τ2 2 1

(3)

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where τ1 and τ2 are the time constants measured at T1 and T2, respectively, that can be written as: ⎛ T + 10 ⎞ Ea = RT ⎜ ⎟ ln Q10 ⎝ 10 ⎠

(4)

ThermoTRP channels present exceptionally high Q10s (higher than 20). With a Q10 of ~26 for TRPV137–39 and ~24 for TRPM840, they far surpass the temperature dependence of the gating processes of other types of ion channels. With some notable exceptions, most channels’ gating processes have Q10 values of about 3.41 For example, in the Shaker K+ channel the inactivation process is highly temperature dependent. The inactivation rate increases with increasing temperature with a Q10 of 7,42 a result that has been interpreted in terms of a temperature-induced stabilization of some structure(s) of the inactivation gate able to bind much more strongly to the channel than the other structures that the peptide can adopt.43 Interestingly, when the inactivating peptide is used to block the channel, with a Q10 of 5, the peptideassociation rate constant is much more temperature sensitive than the dissociation rate constant (Q10 ~1). The slow gating of the ClC0 channel shows a very strong temperature dependence (Q10 ~40), and this result has been interpreted in terms of a coupling of the slow gate with channel subunit interaction.44 In general, a high Q10 for channel gating is compatible with large rearrangements of the protein induced by temperature. We notice here that in the case of TRPM8, the increase in ionic current is mediated by a decrease in temperature so the Q10, as deﬁned in equation (1), would be less than one. This could be wrongly interpreted as a gating process having low temperature sensitivity. To avoid this, the Q10 for channels that respond to cold stimuli has to be deﬁned as the reciprocal of the term on the right in equation (1). This is simply because equation (1) is not general. As we will discuss below, when rates are used to calculate Q10s [e.g., eq. (4)] their values are always higher than one. For thermoTRPs such as TRPV1 and TRPM8, thermodynamic parameters such as the overall changes in enthalpy (H) and entropy (S) associated with channel opening reaction have been obtained considering a two-state model, α ( T ,V ) ⎯⎯⎯ ⎯⎯⎯ →O C← β ( T ,V )

where C and O denote the closed and the open state and the voltage- and temperature-dependent α(T,V) and β(T,V) are the voltage- and temperature-dependent forward (activation) and backward (deactivation) rate constants, respectively. In this type of model, the thermodynamic meaning of the shift of the open probability (Po) vs. voltage curve with temperature is easy to visualize since Po =

1 1 + K −1

(5)

Voltage and Temperature Gating of ThermoTRP Channels

291

where K=

α = e− ΔG / RT β

(6)

in this equation ΔG corresponds to the free energy of the closed-open reaction, given by the relation,45 ΔG = ΔH − T ΔS − zFV

(7)

where z is the overall apparent gating charge, F is Faraday’s constant, and V the applied voltage. The enthalpy and entropy changes associated with the transition between closed and open states have been calculated using the van’t Hoff equation ln K = −

ΔH ΔS + RT R

(8)

and plotting lnK as a function of 1/T, assuming in this case that the term zFV/RT is negligible. Some recent studies have addressed biophysics and thermodynamics of TRPV1, one of the molecular transducers of heat sensation and of its counterpart TRPM8, a cold receptor. Large values for transitional entropy, enthalpy, and Q10 were found in both channels (Table 21.1),37,40 indicating large rearrangements in the channel structure during activation. A remarkable feature of these channels is that their openings are accompanied by very large enthalpic and entropic changes with the net result that the free energy is low.37,40 Thus, the highly temperature-dependent close-open process takes place only because the large enthalpic change, which imparts the channel high-temperature sensitivity, is compensated by a large change in entropy so that the net free energy remains small. However, an increase in temperature induces an increase in ΔG for TRPM8 and a decrease in ΔG for TRPV1. This behavior is a consequence of the opposite signs of the entropic change, which is negative for TRPM8 and positive for TRPV1, indicating that for the cold receptor the closed state has greater entropy than the open state, whereas the opposite is true for TRPV1. Notice that for both TRPV1 and TRPM8 (Table 21.1), ΔH >> zFV because if z = 0.6 and V = 100 mV the term zFV is only 5.8 kcal/mol.

VOLTAGE DEPENDENCE TRPV1 and TRPM8 are weakly voltage dependent, and the apparent number of gating charges (z) calculated from a Boltzmann ﬁt (eq. 9) of the probability of opening (Po) vs. voltage data is 0.6–0.8.40,46,47 Po =

Pomax 1 + exp[− zF (V − V0.5 )/ RT ]

(9)

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Where Pomax is the maximum probability of opening, z the apparent number of gating charges, and V0.5 is the potential at which Po = 0.5. TRPV3 and TRPV4 are also voltage dependent with zs in the range of those found for TRPV1 or TRPM8.20,47,48 The voltage dependence in these channels appears to be intrinsic to the channelforming protein. The voltage sensor of these channels is, however, unknown, and a closer inspection of the predicted S4 segment of these channels reveals the presence of only one basic residue in TRPV1, TRPV3, and TRPV4 and three in TRPM8. It is possible that the weak voltage dependence showed by thermoTRPs is due to the scarcity of positive charges in the S4 domain. In the case of TRPV1 and TRPM8, temperature produces large (>100 mV) leftward shifts of the voltage activation curve upon heating and cooling, respectively.40,46 We notice that when Po = 1/2, equations (5) and (6) imply that K = 1 and ΔG = 0. Inserting this condition in equation (7) gives that the voltage at which the probability is 1/2 is V1/ 2 = ( ΔH − T ΔS )/ zF

(10)

where ΔH and ΔS can be obtained by plotting lnK vs. 1/ T using eq. (8) or V1/2 vs. T using eq. (10). Correa et al.49 realized that the sign from the entropic change during channel opening can be obtained simply by measuring V1/2 at two different temperatures because from equation (10) we have V1/ 2 (T2 ) − V1/ 2 (T1 ) = − ΔS (T2 − T1 )/ zF

(11)

Voets et al.46 have proposed that the temperature-dependent activities of TRPV1 and TRPM8 can be explained, at least in part, by the effects of temperature on the voltage-dependent gating. Because z in thermoTRPs is small, temperature changes promote large shifts of the voltage-activation curves when compared with a channel that has a strong voltage dependence like, for example, Shaker (Figure 21.1A to C). This observation appears to be valid even for more complicated kinetic schemes (Figure 21.1D and E). What makes thermoTRPs special is not their small z, but rather the enormous enthalpic change that imparts to these channels the capability to be gated by temperature as we discuss below. (Note also that Q10 directly relates to the transitional enthalpy, Ea through [eq.] 4.) We further argue that, given the ΔH and ΔS involved in the thermoTRP channels close-open reaction, the most efﬁcient temperature-activated channel would be a voltage-independent one (Figure 21.1C), because temperature would be able to open (or close) the channel at all voltages. From eq. (5), (6), and (7) we have that the change in Po with temperature, assuming that ΔH and ΔS do not change with temperature, is given by the relation: ∂Po ΔH − zFV = ∂T 4 RT 2 cosh 2 ⎛⎜ ΔG ⎞⎟ ⎝ 2 RT ⎠

(12)

Voltage and Temperature Gating of ThermoTRP Channels

293

FIGURE 21.1 Open probabilities as a function of voltage at different temperatures. In a twostate model with ΔH = −86 kcal/mol, ΔS = −297cal/mol°K with (A) z = 2.5 and (B) z = 0.6 and (C) z = 0. For the eight-state allosteric model proposed by Brauchi et al.40 with similar values and an apparent gating charge of (D) zJ = 2.5 and (E) zJ = 0.6.V0.5 vs. temperature plots are shown for each case, with the exception of z = 0, in which the slope diverges.

This function is zero when ΔG → ∞ and tends to (H-zFV)/(4RT2) when ΔG → 0. As thermoTRP channels exhibit relatively small ΔG values during the close-open transition, from equation (12) we see that to ensure large changes in Po with temperature,

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FIGURE 21.1 (Continued).

the only value that has to be large is ΔH. This, in turn, implies that the entropy term has to be large in order to make ΔG small in eq. (7).

PROTEIN DENATURATION, LIPIDS, AND THERMOTRP CHANNELS The most plausible explanation for the effect of temperature on thermoTRP channels considers the existence of a temperature-sensing domain, a “temperature sensor” that would suffer large structural rearrangements upon temperature changes. Protein denaturation is also a highly temperature-dependent process, characterized by large entropic and enthalpic changes, whereas ΔG is relatively small (e.g., Privalov, 1989).50 This is another case in which a highly temperature-dependent process takes place only because the large enthalpic change is compensated by a large entropic change. Heat denaturation is the result of thermal excitation of the numerous disordered conformations accessible to the unfolded polypeptide chain. On the other hand, though the molecular details of cold denaturation remains unclear, it is usually attributed to a weakening of the hydrophobic effect caused by the temperature-dependent structure of bulk water. There are well-described examples of cold denaturation in the same range in which TRPM8 channels are being activated (26–10°C). Phosphoglicerate kinase is denaturated by cold at temperatures ≤30°C,51 and ApoAI denatures at temperatures lower than 25°C.52 In these examples “denaturation” involves loss of tertiary and quaternary interactions, because the subunit interactions are the major target of the process. Among various intermolecular interactions, hydrophobic interactions are strongly temperature dependent as they decrease with temperature and even change signs at sufﬁciently low temperature.53 As hydrophobic interactions are entropically driven, the negative ΔS obtained for TRPM8 during the activation process suggests that there is a net loss of hydrophobic interactions in the closed-to-open transition by temperature. It is important to consider that a protein that shows observable cold denaturation must be characterized by a substantial solvent exposure of apolar groups upon unfolding.54 This suggests that it is possible that in thermoTRP channels, certain specialized regions of the channel suffer structural rearrangements driven by cold, whereas in TRPV1 channels, different rearrangements of this hypothetic “temperature sensor” are driven by heat. Phase changes in the lipid bilayer membrane are well-known phenomena, and this phase transition can have implications in the gating of thermoTRPs induced by

Voltage and Temperature Gating of ThermoTRP Channels

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temperature. Experiments carried out on artiﬁcial lipid bilayer membranes indicate that ionic conductances mediated by the pore formers gramicidin and alamethicin do not show any peculiar effect at the transition temperature (tc) of the phospholipids.55,56 On the other hand, the conductance induced by carriers such as valinomycin is essentially abolished when the temperature is reduced below the lipid phase transition temperature. These results suggest that carriers may freeze out into the membrane/water interface, and channels can be activated even below tc. Whether or not lipid phase changes play a role in thermoTRP channel gating is unknown, and it would be of interest to perform this type of study by incorporating thermoTRP channels into lipid bilayers composed of lipids with well deﬁned tc. Regarding possible effects of membrane lipid composition on thermoTRP channel gating, Liu and colleagues showed that cholesterol depletion did not have a signiﬁcant effect on heat response of the TRPV1 channel but that cholesterol enrichment shifted the half activation temperature in 4°C.37

MULTISTATE NATURE OF THERMOTRPS’ OPENING PROCESS At the single channel level for TRPV1, as temperature increases more openings occur, bursts are prolonged, and quiescent (gaps) periods between bursts are shortened.37 Temperature activates the TRPV1 channel by increasing the burst duration (Q10 ~32) and shortening the gaps (Q10 = 7). Thus, most of the TRPV1 temperature dependence resides in the burst elongation, because the Q10 of the overall single channel activity (Po) was estimated to be 27. Analysis of the channel activity within a burst reveals at least three closed and three open states, the duration of which are only weakly temperature dependent. Capsaicin and pH produce a pattern of single-channel activity similar to that found when temperature is increased.57,58 On the other hand, at the macroscopic current level, for both TRPM8 and TRPV1 the time-dependent changes in conductance due to a membrane depolarization or repolarization occurred with biexponential kinetics.59,40 In the case of TRPV1, temperature affects preferentially the time constant of the slowest component of the current activation (Q10 = 9).46 Temperature only slightly modiﬁes the time constant for the deactivation of TPRV1 channels (Q10 = 1.4). On the other hand, for TRPM8 the time constant of the slowest component of the current deactivation is steeply temperature dependent (Q10 = 4 − 9),40,46 whereas the slowest component of current activation has only a Q10 of 1.4. The deactivation time constant in TRPM8 decreases as temperature decreases, indicating that closing rates are the parameters affected by temperature and explaining, at least in part, why cooling augments TRPM8-induced macroscopic currents, giving rise to an anomalous (<1) Q10. The Q10 for the overall channel gating is ~25 (see Table 21.1), indicating that the temperature-dependent components of the macroscopic channel kinetics have a lower temperature dependence than the overall activation of TRPV1 or TRPM8. The interpretation of the effect of temperature on the activation kinetics can be difﬁcult if the activation pathway is multistep and ramiﬁed, which appears to be the case with thermoTRP channels (see below and Ryu et al.; Brauchi et al.).40,58 In that case, Schoppa and Sigworth60 demonstrated that only for large depolarizations where the backward rate constants are negligible, the time constant calculated from the macroscopic current is a measure of the reciprocal value of the slowest rate

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TABLE 21.1 Thermodynamic Parameters

Protein Na+ channel TRPV1 TRPM8 TRPV3 apoA-2

Q10c

ΔH arrec kcal/mol

1.35–1.53 1.42 ∼1.2 2 —

6.2 7 3 N.D. —

Q10mc

ΔHvH kcal/mol

ΔS cal/mol °K

Thermal Threshold °C

∼2.3 27 24 23 —

−23 150 −112 N.D. 17

−62 470 −384 N.D. 49

≥0 ≥43 ≥25 ≥33 56a

References 67,50 11,37 40 11,19,20 53

c = single channel conductance, arre c = Arrhenius plot activation process enthalpy, mc = macroscopic current, vH = van’t Hoff enthalpy, apoA-2 = human high-density apolipoprotein A-2, a = heat unfolding temperature.

constant in the activation path. In the case of TRPV1, the lower Q10 (~9) of the activation time constant (τa)46 compared to that obtained for the mean burst time (~32) may reﬂect the fact that τa is the inverse of a composite of rate constants with widely different values and temperature sensitivities.

A MINIMUM MODEL THAT EXPLAINS THE CHANGE IN LENGTH OF BURSTS AND GAPS WITH TEMPERATURE The simplest kinetic scheme able to explain single-channel current records in which the opening occurs in bursts separated by gaps is a model containing one open and two closed states: k+ α ⎯⎯ ⎯ → Closed2 ← ⎯⎯ ⎯ → Open Closed1 ← ⎯ ⎯ β

k−

Scheme I

In this model, the bursts consist in a sojourn between the Open and Closed2 states and the gaps in a sojourn in the Closed1 state. To ensure this, the constants k+ and k− have to be much larger than α and β. To predict heat-induced changes in the length of bursts and gaps, α and β have to be highly temperature dependent. According to Eyring’s transition state theory, the kinetic constants are related to the enthalpy and entropy of activation by the following equations

α = νe

⎛ † †⎞ ⎜ ΔH − T ΔS ⎟ α α⎟ ⎜ ⎟ ⎜ RT ⎟ ⎜ ⎠ ⎝

,

β = νe

⎛ † †⎞ ⎜ ΔH − T ΔS ⎟ β β⎟ ⎜ ⎜ ⎟ RT ⎜ ⎟ ⎜⎝ ⎟⎠

(13, 14)

Voltage and Temperature Gating of ThermoTRP Channels

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where ΔH† and ΔS† are the enthalpy and entropy of activation of each rate constant, respectively, and is the prefactor rate that depends on the type of process and here will be assumed to be 106 s-1.61 If the reaction between Closed2 and Open is in equilibrium with respect to Closed1–Closed2 then termination of the burst will be determined by the rate constant β and k1/k-1 and the termination of a gap by α. Introducing the appropriate values (k+ = k-=104 s-1, ΔH†α = 39.5 kcal/mol, ΔS†α = 100 cal/mol°K, ΔH†β = −70 kcal/mol, and ΔS†α = −243 cal/mol°K) in this model, the behavior of the single-channel currents of the TRPV1 channel measured by Liu et al.37 can be reproduced (Figure 21.2A). If the values of ΔH† and ΔS† are exchanged between α and β, we found that the temperature dependence of the gaps is higher than that of the bursts (Figure 21.2B) and it is possible to reproduce qualitatively TRPM8 single channel behavior when the temperature is changed (Figure 21.2C).

AN ALLOSTERIC MODEL FOR THERMOTRP ACTIVATION In TRPM8 channels, we found that a simple way to explain our data was to use an allosteric activation mechanism for both voltage and temperature dependence.40 In this model, we assumed the existence of a “temperature sensor” that has, as does the voltage sensor, resting and activated states (Figure 21.2D). We propose that there is an allosteric linkage between the voltage and temperature sensors or, in other words, neither voltage nor temperature is strictly necessary for channel activation (Figure 21.2D, boxed section). This model can reproduce the steady-state behavior of TRPM8 over a wide range of conditions, with the assumption that activation of voltage sensors and temperature sensors additively affect the energy of the closed-open transitions. The allosteric model successfully accounts for the shift in V0.5 with temperature, noticing that the value of V0.5 tends to saturate at high temperatures and low temperatures (cf. eq. [10]; Figure 21.1D, E, and 21.2F; Voets et al; Brauchi et al.).40,46 This type of model has been very successful in explaining the behavior of another channel activated by two stimuli: the Ca2+- and voltage-activated potassium channel (Slo1; reviewed in Magleby).62

TESTING

THE

ALLOSTERIC MODEL

Figure 21.2C shows the presence of multiple open and closed states in the TRPM8 channel. How these states are connected is not known and requires detailed study of the single channel gating kinetics, in particular the possible correlation between open and closed states.63 The presence of several open and closed states gives, however, support to the model shown in Figure 21.2D. The allosteric model predicts that in the absence of stimuli (i.e., at negative voltages and high temperatures), the channel should be conﬁned to the equilibrium between states C0 and O0 determined by the equilibrium constant L (i.e., the channel can open in the absence of activated voltage or temperature sensors). In this case the probability of opening is given by the equation: L = L (0 )e − zL FV / RT

(15)

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TRP Ion Channel Function in Sensory Transduction

FIGURE 21.2 (A) Relationship between the mean time of gaps and bursts with temperature in the model on scheme I. The values used were k+ = k-= 104 s-1, ΔH†α = 39.5 kcal/mol, ΔS†α = 100 cal/mol°K, ΔH†β = −70 kcal/mol and ΔS†β = −243 cal/mol°K. The formula used for the mean time of bursts and gaps is the same as in Colquhoun and Hawkes.67 (B) The same plot as (A) but the values of ΔH† and ΔS† are exchanged between α and β in a. (C) Upper panel. TRPM8 single-channel recordings at two temperatures: 28°C (low Po) and 20°C (high Po). Recordings were obtained in cell-attached conditions; patches were performed on HEK293 transfected cells. Lower panel. Dwell time frequency distributions for close and open states observed in the recordings above. The best ﬁt to the distributions renders three closed states (28°C: τ1 = 0.57ms; τ2 = 2.95ms; τ3 = 139ms. 20°C: τ1 = 1.1ms; τ2 = 6.3ms; τ3 = 158ms) and two open states (28°C: τ1 = 0.62ms; τ2 = 6.8ms. 20°C: τ1 = 1.25ms; τ2 = 14ms). (D) Allosteric model for activation by voltage and temperature. See text for details. (E) Plot of half-activation voltages (V0.5) against temperature. Symbols are experimental data (mean ± SEM). (F) Current-temperature recordings at holding potentials of −60 and +60 mV. Ramp speed, 0.2ºC/s. Note that voltage changes the temperature threshold for opening.

According to the allosteric model, at sufﬁciently negative voltages voltage sensors should remain in the resting state even when temperature is decreased. Figure 21.2F shows that at negative voltages (−60 mV), TRPM8-induced current is highly temperature sensitive albeit with a lower temperature threshold when compared to the data

Voltage and Temperature Gating of ThermoTRP Channels

299

obtained at +60 mV. Thus, as the allosteric model predicts, a decrease in temperature can activate TRPM8 channels when all voltage sensors are in the resting state. Because the channel is a homotetramer, it is likely to contain four voltage sensors and the same number of temperature-sensing structures, and this clearly will increase the number of states to at least 50. The situation, however, is not desperate, because the number of parameters can be reduced (and determined) working under extreme voltages or temperatures and measuring and characterizing single-channel and macroscopic currents. It is necessary to understand thermoTRPs’ gating mechanisms in terms of models that include the large number of open and closed states (e.g., Liu et al.).37 Allosteric models offer a natural explanation to the single-channel data, assuming that the channel has multiple heat-sensitive sites distributed over the subunits. Activation of one site brings about a short burst of activity. As temperature is increased, more sites become active, and if they interact allosterically, this interaction will produce elongation of the bursts as observed experimentally in TRPV1.37 Allosteric models have been successful in explaining the workings of other channels gated by several stimuli (e. g., Horrigan and Aldrich; Orio and Latorre)64,65 and can be particularly useful to describe the complex regulation of TRPM8 by agonists such as PIP2 and cooling agents and that of TRPV1 by capsaicin and pH.

REFERENCES 1. Clapham, D.E., TRP channels as cellular sensors, Nature, 426, 517, 2003. 2. Montell, C., An end insight to a long TRP, Neuron, 30(1), 3, 2001. 3. Montell, C., Birnbaumer, L., and Flockerzi, V., The TRP channels, a remarkably functional family, Cell, 108, 595, 2002. 4. Moran, M.M., Xu, H., and Clapham D.E., TRP ion channels in the nervous system, Curr. Opin. Neurobiol., 14(3), 362, 2004. 5. Voets, T., Talavera, K., Owsianik, G., and Nilius, B., Sensing with TRP channels, Nature Chem. Biol., 1(2), 85, 2005. 6. Kedei, N. et al., Analysis of quaternary structure of vanilloid receptor 1, J. Biol. Chem., 276(30), 28613, 2001. 7. Hellwig, N. et al., Homo- and heteromeric assembly of TRPV channel subunits, J. Cell Sci., 118(5), 917, 2005. 8. Tominaga, M. and Caterina, M.J., Thermosensation and pain, J. Neurobiol., 61(1), 3, 2004. 9. Benham, C.D., Gunthorpe, M.J., and Davis, J.B., TRPV channels as temperature sensors, Cell Calcium, 33, 479, 2003. 10. McKemy, D.D. How cold is it? TRPM8 and TRPA1 in the molecular logic of cold sensation, Mol. Pain, 1(1), 16, 2005. 11. Caterina, M.J. et al. The capsaicin receptor: a heat-activated ion channel in the pain pathway, Nature, 389, 816, 1997. 12. Tominaga, M. et al. The cloned capsaicin receptor integrates multiple pain-producing stimuli, Neuron, 21, 531, 1998. 13. Macpherson, L.J. et al. The pungency of garlic: activation of TRPA1 and TRPV1 in response to allicin, Curr. Biol, 15, 929, 2005. 14. Caterina, M.J. et al. Impaired nociception and pain sensation in mice lacking the capsaicin receptor, Science, 288, 306, 2000.

300

TRP Ion Channel Function in Sensory Transduction

15. Caterina, M.J. et al. A capsaicin-receptor homologue with a high threshold for noxious heat, Nature, 398, 436, 1999. 16. Ma, H.T. et al., Assessment of the role of the inositol 1,4,5-trisphosphate receptor in the activation of transient receptor potential channels and store-operated Ca2+ entry channels, J. Biol. Chem., 276(22), 18888, 2001. 17. Ahluwalia, J., Rang, H., and Nagy, I., The putative role of vanilloid receptor–like protein-1 in mediating high-threshold noxious heat-sensitivity in rat cultured primary sensory neurons, Eur. J. Neurosci., 16(8), 1483, 2002. 18. Lewinter, R.D. et al., Immunoreactive TRPV-2 (VRL-1), a capsaicin receptor homolog, in the spinal cord of the rat, J. Comp. Neurol., 470(4), 400, 2004. 19. Smith, G.D. et al., TRPV3 is a temperature-sensitive vanilloid receptor–like protein, Nature, 418, 186, 2002. 20. Xu, H. et al., TRPV3 is a calcium-permeable temperature-sensitive cation channel, Nature, 418, 181, 2002. 21. Peier, A.M. et al., A heat-sensitive TRP channel expressed in keratinocytes, Science, 296(5575), 2046, 2002a. 22. Chung, M.K., Guler, A.D., and Caterina, M.J., Biphasic currents evoked by chemical or thermal activation of the heat-gated ion channel, TRPV3, J. Biol. Chem., 280(16), 15928, 2005. 23. Moqrich, A. et al., Impaired thermosensation in mice lacking TRPV3, a heat and camphor sensor in the skin, Science, 307(5714), 1468, 2005. 24. Liedtke, W. et al., Vanilloid receptor–related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor, Cell, 103(3), 525, 2000. 25. Strotmann, R. et al., OTRPC4, a nonselective cation channel that confers sensitivity to extracellular osmolarity, Nat. Cell Biol., 2(10), 695, 2000. 26. Güler, A.D. et al., Heat-evoked activation of the ion channel, TRPV4, J. Neurosci., 22(15), 6408, 2002. 27. Watanabe, H. et al., Heat-evoked activation of TRPV4 channels in a HEK293 cell expression system and in native mouse aorta endothelial cells, J. Biol. Chem., 277(49), 47044, 2002. 28. Chung, M.K. et al., 2-aminoethoxydiphenyl borate activates and sensitizes the heatgated ion channel TRPV3, J. Neurosci., 24(22), 5177, 2004a. 29. Chung, M.K. et al., TRPV3 and TRPV4 mediate warmth-evoked currents in primary mouse keratinocytes, J. Biol. Chem., 279(20), 21569, 2004b. 30. McKemy, D.D., Neuhausser, W.M., and Julius, D., Identiﬁcation of a cold receptor reveals a general role for TRP channels in thermosensation, Nature, 416(6876), 52, 2002. 31. Peier, A.M. et al., A TRP channel that senses cold stimuli and menthol, Cell, 108(5), 705, 2002b. 32. Reid, G., ThermoTRP channels and cold sensing: what are they really up to? Pﬂügers Arch. 2005 June 17. 33. Story, G.M. et al., ANTKTM1, a TRP-like channel expressed in nociceptive neurons, is activated by cold temperatures, Cell, 112, 819, 2003. 34. Jordt, S.E. et al., Mustard oils and cannabinoids excite sensory nerve ﬁbres through the TRP channel ANTKTM1, Nature, 427, 260, 2004. 35. Bandell, M. et al., Noxious cold ion channel TRPA1 is activated by pungent compounds and bradykinin, Neuron, 41(6), 849, 2004. 36. Babes, A., Zorzon, D., and Reid, G., Two populations of cold-sensitive neurons in rat dorsal root ganglia and their modulation by nerve growth factor, Eur. J. Neurosci., 20(9), 2276, 2004.

Voltage and Temperature Gating of ThermoTRP Channels

301

37. Liu, B., Hui, K., and Qin, F., Thermodynamics of heat activation of single capsaicin ion channels VR1, Biophys. J., 85, 2988, 2003. 38. Nagy, I. and Rang, H.P. Similarities and differences between the responses of rat sensory neurons to noxious heat and capsaicin, J. Neurosci., 19(24), 10647, 1999. 39. Vlachova, V. et al., Functional role of C-terminal cytoplasmic tail of rat vanilloid receptor 1, J. Neurosci., 23(4), 1340, 2003. 40. Brauchi, S., Orio, P., and Latorre, R., Clues to understanding cold sensation: thermodynamics and electrophysiological analysis of the cold receptor TRPM8, Proc. Natl. Acad. Sci. USA, 101(43), 15494, 2004. 41. Hille, B. Ionic channels of excitable membranes, Sinauer Associates, Sunderland, MA, 2001. 42. Nobile, M. et al., Fast inactivation of Shaker K+ channels is highly temperature dependent, Exp. Brain Res., 114(1), 138, 1997. 43. Murrell-Lagnado, R.D. and Aldrich, R.W., Energetics of Shaker K channels block by inactivation peptides, J. Gen. Physiol., 102(6), 977, 1993. 44. Pusch, M., Ludewig, U., and Jentsch, T.J., Temperature dependence of fast and slow gating relaxations of CIC-O chloride channels, J. Gen. Physiol., 109(1), 105, 1997. 45. Lecar, H., Ehrenstein, G., and Latorre, R., Mechanism for channel gating in excitable bilayers, Ann. NY Acad. Sci., 264, 304, 1975. 46. Voets, T. et al., The principle of temperature-dependent gating in cold- and heatsensitive TRP channels, Nature, 430(7001), 748, 2004. 47. Nilius, B. et al., Gating of TRP channels: a voltage connection? J. Physiol., 567(1), 35, 2005. 48. Chung, M.K., Guler, A.D., and Caterina, M.J., Biphasic currents evoked by chemical or thermal activation of the heat-gated ion channel, TRPV3, J. Biol. Chem., 280(16), 15928, 2005. 49. Correa, A.M., Bezanilla, F., and Latorre, R., Gating kinetics of batrachotoxinmodiﬁed Na+ channels in the squid giant axon. Voltage and temperature effects. Biophys. J., 61(5), 1332, 1992. 50. Privalov, P.L., Thermodynamic problems of protein structure, Annu. Rev. Biophys. Biophys. Chem., 18, 47, 1989. 51. Griko, Y.V., Venyaminov, S.Y., and Privalov, P.L., Heat and cold denaturation of phosphoglycerate kinase (interaction of domains), FEBS Lett., 244(2), 276, 1989. 52. Gursky, O. and Atkinson, D. Thermal unfolding of human high-density apolipoprotein A-1: implications for a lipid-free molten globular state, Proc. Natl. Acad. Sci. USA, 93(7), 2991, 1996. 53. Privalov, P.L. and Gill, S.J., Stability of protein structure and hydrophobic interaction, Adv. Protein Chem., 39, 191, 1988. 54. Privalov, P.L., Cold denaturation of proteins, Crit. Rev. Biochem. Mol. Biol., 25(4), 281, 1990. 55. Krasne, S., Eisenman, G., and Szabo, G., Freezing and melting of lipid bilayers and the mode of action of nonactin, valinomycin, and gramicidin, Science, 174(7), 412, 1971. 56. Boheim, G., Hanke, W., and Eibl, H., Lipid phase transition in planar bilayer membrane and its effect on carrier- and pore-mediated ion transport, PNAS, 77(6), 3403, 1980. 57. Hui, K., Liu, B., and Qin, F., Capsaicin activation of the pain receptor, VR1: multiple open states from both partial and full binding, Biophys. J., 84, 2957, 2003.

302

TRP Ion Channel Function in Sensory Transduction

58. Ryu, S., Liu, B., and Qin, F., Low pH potentiates both capsaicin binding and channel gating of VR1 receptors, J. Gen. Physiol., 122, 45, 2003. 59. Ahern, G.P. and Premkumar, L.S., Voltage-dependent priming of rat vanilloid receptor: effects of agonist and protein kinase C activation, J. Physiol., 545(2), 441, 2002. 60. Schoppa, N.E. and Sigworth, F.J., Activation of Shaker potassium channels. III. An activation gating model for wild-type and V2 mutant channels. J. Gen. Physiol., 111(2), 313, 1998. 61. Yang, W.Y. and Gruebele, M., Folding at the speed limit, Nature, 423(6936), 193, 2003. 62. Magleby, K.L., Gating mechanism of BK (Slo1) channels: so near, yet so far. J. Gen. Physiol., 121(2), 81, 2003. 63. McManus, O.B., Blatz, A.L., and Magleby, K.L., Inverse relationship of the durations of adjacent open and shut intervals for C1 and K channels. Nature, 317(6038), 625, 1985. 64. Horrigan, F.T. and Aldrich, R.W., Coupling between voltage sensor activation, Ca2+ binding and channel opening in large conductance (BK) potassium channels, J. Gen. Physiol., 120(3), 267, 2002. 65. Orio, P. and Latorre, R., Differential effects of beta 1 and beta 2 subunits on BK channel activity, J. Gen. Physiol., 125(4), 395, 2005. 66. Horn, R., Vandenberg, C.A., and Lange, K., Statistical analysis of single sodium channels. Effects of N-bromoacetamide, Biophys. J., 45(1), 323, 1984. 67. Colquhoun, D. and Hawkes, A.G. On the stochastic properties of single ion channels, Proc. R. Soc. Lond., 211, 205, 1981.

22

TRPV Channels’ Function in Osmo- and Mechanotransduction Wolfgang B. Liedtke Duke University

CONTENTS Abstract ..................................................................................................................303 Introduction: The TRPV Subfamily ......................................................................304 Heterologous Cellular Expression Systems ..........................................................304 Osmo- and MechanoTRP TRPV1: Animal Findings ...........................................305 Osmo- and MechanoTRP TRPV2: Tissue Culture Data ......................................306 Osmo- and MechanoTRP TRPV4: Tissue Culture and Animal Data ..................307 C.Elegans TRPV Channels and Mechano-, Osmosensation.................................310 Cloning of the osm-9 Gene, a Founding Member of the trpv Gene Family ............................................................................310 TRPV4 Expression in ASH Rescues osm-9 Mechanical and Osmotic Deﬁcits .................................................................................311 Outlook for Future Research on TRP Channels ...................................................313 Acknowledgment ...................................................................................................313 References..............................................................................................................314

ABSTRACT In signal transduction of metazoan cells, transient receptor potential (TRP) ion channels have been identiﬁed to respond to diverse external and internal stimuli, among them osmotic and mechanical stimuli. This chapter summarizes ﬁndings on the TRPV subfamily, both its vertebrate and invertebrate members, with a focus on TRPV4. Of the six mammalian TRPV channels, TRPV1, 2, and 4 were demonstrated to function in transduction of osmotic and mechanical stimuli. Invertebrate TRPV channels, ﬁve in C. elegans and two in Drosophila, have been shown to play a role in mechanosensation, such as hearing and proprioception in Drosophila and nose touch in C. elegans, and in the response to tonicity in C. elegans. TRPV4 has been found to function in cellular as well as systemic osmotic homeostasis in vertebrates. In a striking example of evolutionary conservation of function, mammalian TRPV4

303

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has been found to rescue mechano- and osmosensory, not olfactory, deﬁcits of the TRPV mutant line osm-9 in C. elegans, despite not more than 25 percent orthology of the respective amino acid sequences.

INTRODUCTION: THE TRPV SUBFAMILY Within the TRP superfamily of ion channels [21], the TRPV subfamily came into existence and immediately gained public notoriety in 1997 [14,19], when its founding members, TRPV1 in mammals and OSM-9 in C. elegans, were ﬁrst reported. TRPV1 was identiﬁed by an expression cloning strategy [14]. OSM-9 was identiﬁed through genetic screening for worms’ defects in osmotic avoidance [19]. TRPV2, -V3, and -V4 were identiﬁed by a candidate gene approach, respectively [13,36,43,56,60,63,74,76]. The latter strategy also led to the identiﬁcation of four additional C. elegans ocr genes [66] and two Drosophila trpv genes, Nanchung (NAN) and Inactive (IAV) [28,38]. The TRPV channels can be subgrouped into four branches by sequence comparison. One branch includes four members of mammalian TRPVs (TRPV1, -V2, -V3, and -V4); in vitro whole-cell recording showed that they respond to temperatures higher than 42, 52, 31, and 27°C, respectively, suggesting that they are involved in thermosensation, hence the term “thermoTRPs” (reviewed in references 10, 12, 18, 55, and 69, and many other meritorious papers). One invertebrate branch includes C. elegans OSM-9 and Drosophila IAV; the other branch is composed of OCR-1 to -4 in C. elegans and Drosophila NAN. This chapter focuses on the role of mammalian (in the ﬁrst place) and also invertebrate (C. elegans) TRPV channels in signal transduction in response to osmotic and mechanical stimuli, in particular on TRPV4. These osmo- and mechanoTRPs [45] are TRPV1, -2, -4, OSM-9, OCR-2, NAN, and IAV. Other TRPV channels might join this functional group within the TRP superfamily, which certainly also contains non-TRPV channels such as TRPA1 [20,52] or NompC [72]. The following issues are addressed in this chapter. Do TRPV ion channels function in sensing and transduction of osmotic and mechanical stimuli? Which molecular mechanisms are at play? Are the responses to either stimulus linked, possibly via transduction of membrane tension? When answering these questions, one has to bear in mind that (1) the ﬁeld of TRPV ion channels is young (less than a decade) [14,19] and that (2) the methodological arsenal to answer the above questions for TRPVs has its limitations.

HETEROLOGOUS CELLULAR EXPRESSION SYSTEMS Before the TRPV osmo- and mechanoTRPs receive further consideration, here is a short discourse regarding limitations of heterologous cellular expression systems, which have helped elucidate TRP channels’ function, yet the following qualiﬁers will have to be realized. For the ﬁeld of ion channel physiology, heterologous cellular expression systems have allowed particularly rewarding studies for voltage-gated channels and also for ligand-gated channels such as the nicotinic acetylcholine-receptor, GABA-ergic

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channels, and NMDA receptors. It is perhaps slightly underappreciated that this concept cannot be transferred seamlessly to the investigation of channels that respond to osmotic and mechanical stimuli. Nonspeciﬁc effects may be caused by purely physical effects of these stimuli on the cells, for example, mechanical stimulation. First, a latency has to be determined in order to be able to differentiate direct mechanical activation of the channel (i.e., mechanotransduction happens only by activation of the channel without other signaling molecules directly involved in this signaling) versus an indirect activation (i.e., mechanotransductory channel activation downstream). For a direct response, a latency shorter than one millisecond is required [33–35]. With currently available technology, this means that only patch-clamp recordings satisfy this need. One problem is how to apply the mechanical stimulus without disturbing the recording. With respect to tonicity, the precise beginning of the osmotic stimulus cannot be determined. When applying the osmotic stimulus by streaming bath solution, one has to realize that a mechanical stimulus is coapplied, namely ﬂow, which will affect the cell by exerting shear stress. Also, osmotic and mechanical stimuli as activators of channels are distinctly different from speciﬁc activators/ligands (e.g., GABA, NMDA). Most cells will harbor an innate response to primal biophysical stimuli such as tonicity and touch. This means that heterologously expressing a certain ion channel in this context is supplementing a preexisting signaling apparatus by one more molecule. It is quite obvious, on the other hand, that the situation is different for, for example, the response of an epithelial cell or ﬁbroblast to a nervous system–speciﬁc ligand/activator such as GABA, glycine, or NMDA.

OSMO- AND MECHANOTRP TRPV1: ANIMAL FINDINGS In heterologous cellular expression systems, there have not been reports on mechanotransduction by TRPV1. Genetically engineered trpv1−/− mice, which have previously been shown to be devoid of thermal hyperalgesia following inﬂammation [11,23], also displayed an altered response of their bladders to stretching [9]. TRPV1 could be localized to sensory and autonomous ganglia neurons and also to urethelial cells lining the pyelon, ureter, and bladder. When bladder and urothel epithelial cells were maintained in primary tissue cultures, their responses to mechanical stretch were signiﬁcantly different from wild-type cultures. Speciﬁcally, TRPV1+ bladders secreted ATP upon mechanical stretch, which, in turn, is known to activate nerve ﬁbers in the bladder submucosa. This response to mechanical stimulation was greatly reduced in bladders excised from trpv1−/− mice. It appears likely that this mechanism, operative in mice, also plays a role in human bladder epithelia. Intravesical installation of TRPV1 activators is used to treat hyperactive bladder syndromes in spinal cord disease, although the exact effect and mechanism of action of TRPV1 agonists is not clear [4,26,40,62]. Another instance of an altered response to mechanical stimuli in trpv1−/− mice relates to the response of the jejunum to mechanical stretch [58]. Afferent jejunal nerve ﬁbers were found to respond with decreased frequency of discharge in trpv1−/− mice than in wild-type mice [22]. In humans, TRPV1 positive nerve ﬁbers in the rectum were signiﬁcantly increased in patients suffering from

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fecal urgency, a pathologic rectal hypersensitivity in response to mechanical distension [16]. Expression of TRPV1+ nerve ﬁbers in rectal biopsy samples from these patients correlated with a lower threshold to mechanical stretch; in addition, the occurrence of TRPV1+ ﬁbers was also correlated with a dysaesthesia of a burning quality. Another recent study focused on possible mechanisms of signal transduction in response to mechanical stimuli in blood vessels [59]. Elevation of intraluminal mechanical pressure in mesenterial arteries was reported to be associated with generation of 20-hydroxyeicosatetraenoic acid, which, in turn, activated TRPV1 on C-ﬁbers leading to nerve depolarization and vasoactive neuropeptide release. With respect to nociception, using trpv1−/− mice, trpv1 was shown to be involved in inﬂammatory thermal hyperalgesia, but not in inﬂammatory mechanical hyperalgesia [10,32]. However, a speciﬁc and potent blocker of TRPV1 was found to reduce mechanical hyperalgesia in rats [22,57]. These latter results appear in contrast to the lack of difference between trpv1−/− and wild-type mice. Either this discrepancy is due to a species difference between mice and rats pertaining to signal transduction by TRPV1 in inﬂammation-induced mechanical hyperalgesia, or it may be due to the different mechanisms that affect signaling in a trpv1 general knockout (with likely compensatory gene regulation) versus a speciﬁc temporal pharmacological blocking of the TRPV1 ion channel protein that most likely participates in a signaling multiplex. Very recently, reporting a spectacular ﬁnding, Sharif Naeini et al. reported that trpv1−/− mice failed to express an N-terminal variant of the trpv1 gene in magnocellular neurons of the supraoptic and paraventricular nucleus of the hypothalamus [51]. These neurons are known to secrete vasopressin, and the trpv1−/− mice were found to have a profound impairment of ADH secretion in response to systemic hypertonic stimuli, and their magnocellular neurons did not show an appropriate electrical response to hypertonicity. Bourque and colleagues [51] conclude that this trpv1 N-terminal variant, which could not be identiﬁed at the molecular level, is likely involved as (part of) a tonicity sensor of intrinsically osmosensitive magnocellular neurons. Moreover, Ciura and Bourque [81] report that this splice variant is also expressed in the osmotically sensitive circumventricular organ OVLT, and that OVLT-dissociated neurons from the trpv1−/− mice are defective in their response to hypertonic stimuli.

OSMO- AND MECHANOTRP TRPV2: TISSUE CULTURE DATA With respect to the TRPV2 ion channel, we are still awaiting the report on an eventual phenotype of trpv2−/− mice. In heterologous cellular systems, TRPV2 was initially described as a temperature-gated ionotropic receptor for stimuli >52°C [13]. Recently, TRPV2 was also shown to respond to hypotonicity and mechanical stimulation [49]. Arterial smooth muscle cells from various arteries expressed TRPV2. These myocytes responded to hypotonic stimulation with calcium inﬂux. This activation could be diminished by speciﬁc downregulation of TRPV2 protein by an antisense strategy. Heterologously expressed TRPV2 in CHO cells displayed a similar response to hypotonicity. These cells were also subjected to stretch by applying negative pressure to the patch pipette and by stretching the cell membrane on a

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mechanical stimulator. Both maneuvers led to Ca2+ inﬂux that depended on heterologous TRPV2 expression. TRPV3 has not (yet) been characterized as an osmo- and mechanoTRP, either in heterologous systems or in live animals or human studies. The same is true for TRPV5 and TRPV6.

OSMO- AND MECHANOTRP TRPV4: TISSUE CULTURE AND ANIMAL DATA CHO cells responded to hypotonic solution when they were (stably) transfected with TRPV4 [43]. HEK293 T cells, when maintained by the same authors, were found to express trpv4 cDNA, which was cloned from these cells (genbank AF263523). However, trpv4 cDNA was not found in other batches of HEK293 T cells, so that this cell line was used as a heterologous expression vehicle by other groups [63,74]. When comparing the two settings, it was obvious that the single-channel conductance was not at all similar [43,63]. This perhaps underscores the relevance of gene expression in heterologous cellular systems for the functioning of TRPV4 in response to a basic biophysical stimulus. Also, the sensitivity of TRPV4 could be tuned by warming of the media. Peak sensitivity of gating in response to hypotonicity was recorded at core body temperature of the respective organism, and TRPV4 channels from both birds (core body temp. 40°C) and mammals (37°C) were compared, again in CHO cells [43]. Similar results were found in another investigation with expression of mammalian TRPV4 in HEK293 T cells [27]. In this investigation, the cells were mechanically stretched at isotonicity. At room temperature, there was no response upon mechanical stretch; however, at 37°C the isotonic response to stretch resulted in a very strong calcium inﬂux. In two other investigations, TRPV4 was found to be responsive to changes in temperature [31,73]. Temperature change was accomplished by heating the streaming bath solution (see above comments on ﬂow as a mechanical stimulus). Gating was augmented when hypotonic solution was used as a streaming bath. In one investigation, temperature stimulation could not activate the TRPV4 channel in cell-detached inside-out patches [73]. With respect to the gating mechanism of TRPV4 in response to hypotonicity, two recent papers report conﬂicting results on phosphorylation sites of TRPV4 that are necessary for the response to hypotonicity. One paper reported that TRPV4 was tyrosine-phosphorylated in HEK293 T cells and in distal convoluted tubule cells from mouse kidneys [65,77]. Tyrosine phosphorylation was sensitive to speciﬁc inhibition of the Src family tyrosine kinases. The Lyn tyrosine kinase was found to coimmunoprecipitate with TRPV4 and to feature a critical role in phosphorylation of TRPV4 (Y253). A point mutation of Y253 reduced hypotonicity-induced gating. On the other hand, in another investigation, in HEK293 T cells, hypotonicity activated TRPV4 by phospholipaseA2–mediated formation of arachidonic acid via a cytochrome P450 epoxygenase pathway [71]. In HEK cells, this signaling mechanism did not apply for TRPV4 gating by increased temperature or by the nonphosphorylating phorbol ester 4-alpha PDD. This latter activation mechanism was reported to depend on phosphorylation of Y555. However, the authors of this study could not replicate the aforementioned

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ﬁnding of tyrosine kinase phosphorylation of Y253 of TRPV4 as critical for hypotonicity-induced gating. Why this discrepancy? It reiterates the pivotal role of the host cell in heterologous expression experiments. In another recent paper, the ciliary beat frequency of ciliated cells was shown to be inﬂuenced by TRPV4 gating [3]. In primary ciliated cells, and also in heterologously transfected HeLa cells, TRPV4 could be activated (mechanically) by exposing the cells to hyperviscous, isotonic media. Another recent focus in the ﬁeld of TRP ion channels is intracellular trafﬁcking, posttranslational modiﬁcation, and subsequent functional modulation. For TRPV4, it was found in heterologous cells (HEK293T) that N-glycosylation between the ﬁfth transmembrane domain and pore loop (position 651) decreases osmotic activation via decreased plasma membrane insertion [75]. Interestingly, N-glycosylation between the ﬁrst and second transmembrane domains appears to have the same effect on TRPV5, and the anti-aging hormone klotho functions as beta-glucuronidase and subsequently activates TRPV5 [17]. In the kidney (and via systemic klotho possibly elsewhere), klotho could have a similar effect on TRPV4 and function as an ampliﬁer of tonicity-mediated signaling. TRPV4 also has played a role in maintaining cellular osmotic homeostasis. One particular cellular defense mechanism of cellular osmotic homeostasis is regulatory volume change, namely regulatory volume decrease (RVD) in response to hypotonicity and regulatory volume increase (RVI) in response to hypertonicity. In a recent paper, Bereiter-Hahn’s group reported that CHO tissue culture cells have a poor RVD, which, after transfection with TRPV4, improves in a striking manner [8]. In another study, Valverde’s group published that TRPV4 mediates cell swelling– induced Ca2+ inﬂux into bronchial epithelial cells that triggers RVD via Ca2+dependent potassium channels [6]. This cell-swelling response was not operational in cystic ﬁbrosis (CFTR) bronchial epithelia, where, on the other hand, TRPV4 could be activated by 4-alpha-PDD, leading to Ca2+ inﬂux. Thus, in CFTR bronchial epithelia, RVD could not be elicited by hypotonicity but by 4-alpha-PDD. In yet another investigation, Ambudkar and colleagues found a concerted interaction of aquaporin-5 (AQP-5) with TRPV4 in hypotonic swelling–induced RVD of salivary gland epithelia [42]. These exciting ﬁndings elucidate mechanisms that maintain function of secretory epithelia (such as salivary, tear, sweat, airway, and intestinal glands) that underlie watery secretion based on a concerted interaction of TRPV4 with AQP-5 (for another investigation pertaining to this topic see reference 82). In trpv4−/− mice, the response to noxious mechanical stimulation is diminished [44,64]. In the absence of TRPV4, the threshold to noxious mechanical stimulation was signiﬁcantly elevated. This result was obtained using two standard tests, the Randall-Sellito test, which applies mechanical pressure by squeezing the paw, and an automatized von Frey test, which applies mechanical stimulation from underneath the hindpaw, leading to withdrawal [44]. In mice, paw withdrawal in response to a noxious temperature was not different between trpv4−/− mice and wild-type mice [44,64]. However, a more detailed testing of abnormalities in response to thermal stimuli revealed an abnormal inﬂammatory hyperalgesia in trpv4−/− mice and an

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altered behavior in a thermal gradient [41,68]. When rats were sensitized with taxol, their sensitivity to noxious mechanical stimuli was strikingly lowered as a result of the taxol-induced neuropathy [24,25]. When these rats were treated intrathecally with TRPV4-speciﬁc antisense oligonucleotides, taxol-induced mechanical hypersensitivity was eliminated [1]. This clearly suggests a role for TRPV4 in mediating hyperalgesia in response to mechanical stimuli in an animal model for neuropathic pain. Last, but not least, with respect to mechanotransduction in trpv4−/− mice, they do not show signs of inner ear dysfunction including deafness [44], which has to be viewed against the expression pattern of trpv4 in the inner ear [43,50]. trpv4 mRNA could be demonstrated in the secretory epithelia of the stria vascularis/ tegmentum vasculare and in neurosensory inner ear hair cells of both rodents and birds. This negative ﬁnding in vivo does not exclude, however, a role for TRPV4 in inner ear function. trpv4−/− mice, when stressed with systemic hypertonicity, did not counterregulate their systemic tonicity as efﬁciently as wild-type littermates [44]. Their drinking was reduced, and systemic tonicity was signiﬁcantly higher. Continuous infusion of the ADH analogue dDAVP led to systemic hypotonicity, whereas renal water readsorption capacity was not altered between genotypes. Antidiuretic hormone synthesis in response to osmotic stimulation was reduced in trpv4−/− mice. Hypertonic stress led to reduced expression of c-FOS+ cells in the sensory circumventricular organ, OVLT, indicative of impaired osmotic activation. These ﬁndings in trpv4−/− mice point toward a deﬁcit of osmotic sensing in the central nervous system. Thus, TRPV4 is necessary for maintaining systemic osmotic equilibrium in mammals. It is conceivable that TRPV4 acts as an osmotic sensor in the CNS. The impaired osmotic regulation in trpv4−/− mice reported in the author’s paper differs from that published in another report. While our experiments showed that trpv4−/− mice secrete lower amounts of ADH in response to hypertonic stimuli, the results from Mizuno et al. [48] suggest that there is an increased ADH response to water deprivation and subsequent systemic administration of propylene glycol. The reasons for this discrepancy are not obvious. In our study, a blunted ADH response and diminished cFOS response in the OVLT in trpv4−/− mice upon systemic hypertonicity suggest, as one possibility, an activation of TRPV4+ sensory cells in the OVLT by hypertonicity. This consideration, in contradiction with results from heterologous expression systems, is important and will be extended below. However, together with the above ﬁndings of the Bourque group on abnormal osmotic regulation in trpv1−/− mice, the fascinating possibility is that TRP(V) channels might interact to form critical transduction multiplex in the regulation of systemic tonicity. In other words, is Verney’s osmoreceptor made up of heteromers derived from the trpv1/4 genes? In a recent publication, Alessandri-Haber et al. demonstrate that hypertonic subcutaneous solution leads to pain-related behavior in wild-type mice, which is not present in trpv4−/− mice [2]. When sensitizing nociceptors with prostaglandin E2, the pain-related responses to hypertonic stimulation became more frequent and were greatly reduced in trpv4−/− mice. These in vivo data could not be recapitulated in acutely dissociated DRG neurons upon stimulation with hypertonicity

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and subsequent calcium imaging; there was a discernible rise in intracellular calcium, yet genotypes did not differ. As for hypotonic stimulation, this did not elicit pain-related behavior in mice, which could only be evoked after presensitization with PGE2. And again, this behavior was strikingly reduced in the absence of trpv4. Contrary to hypertonicity, hypotonicity led to an increase of intracellular calcium in primary DRG neurons, yet intracellular calcium was signiﬁcantly reduced in the absence of TRPV4. Taken together, this investigation indicates differences in the response of mice to noxious tonicity stimuli depending on the presence or absence of TRPV4. Yet at the level of a critical transducer cell, namely the DRG sensory neuron in acutely dissociated culture, only hypotonicity led to a rise of intracellular calcium that depended on the presence of TRPV4. Perhaps other closely associated cells, such as epidermal keratinocytes or Schwann cells, assume a critical role in the transduction of hypertonic noxious stimuli or, as a nonmutually exclusive possibility, the in vivo situs where the sensory neuron extends a process that measures several hundred or thousand times the diameter of the soma, is not reﬂected faithfully by the reductionist model of the primary cultured DRG neuron.

C. ELEGANS TRPV CHANNELS AND MECHANO-, OSMOSENSATION CLONING

OF THE OSM-9

OF THE TRPV

GENE,

A

FOUNDING MEMBER

GENE FAMILY

As mentioned in the introduction, the osm-9 mutant was ﬁrst reported in 1997 [19]. The forwards genetics screen in C. elegans consisted of a conﬁnement assay with a high-molar osmotically active substance. osm-9 mutants did not respect this barrier, and the mutated gene was found to be a TRP channel. On closer analysis, osm-9 mutants did not respond to aversive tonicity stimuli, they did not respond to mechanical tapping of their “noses,” and they did not respond to (aversive) odorants. The OSM-9 channel protein was expressed in amphid sensory neurons, the worms’ cellular substrate of exteroceptive sensing of noxious chemical, osmotic, and mechanical stimuli. The OSM-9 channel was expressed in the sensory cilia of the AWC and ASH amphid sensory neurons. Bilateral laser ablation of the ASH neuron has led to a deﬁcit in avoidance of noxious osmotic, nose touch, and olfactory stimuli [37], hence the term “nociceptive” neuron [7]. The OSM-9 protein could not, however, be expressed in heterologous cellular expression systems, and explant cultures of amphid sensory neurons were not viable. Next, also by the Bargmann laboratory, four additional TRPV channels from C. elegans were cloned, named OCR-1 to -4 [66]. Of these four channels, only the OCR2 channel was expressed in ASH. The ocr-2 mutant phenotype was virtually identical to the osm-9 phenotype with respect to worm “nociception,” and there was genetic evidence that the two channels interacted. When expressing the mammalian capsaicin receptor TRPV1 in the ASH sensory neurons, neither osm-9 nor ocr-2 mutants could

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be rescued for any of their deﬁcits, but osm-9 ash::trpv1 transgenic worms displayed a strong avoidance response to capsaicin, which normal worms virtually do not show.

TRPV4 EXPRESSION IN ASH RESCUES AND OSMOTIC DEFICITS

OSM-9

MECHANICAL

Next, TRPV4 was transgenically targeted to ASH of osm-9 mutants. Surprisingly, this rescued osm-9 mutants’ defects in avoidance of hyperosmotic noxious stimuli and nose touch [46], not odorant avoidance of osm-9, suggesting that this speciﬁc function of TRPV channels differs between vertebrates and invertebrates. This basic ﬁnding of the rescue experiments in osm-9 ash::trpv4 worms has implications for mechanisms of signal transduction in the ASH neuron (see Figure 22.1). TRPV4 appeared to be integrated into the normal ASH sensory neuron signaling apparatus, because the transgene failed to rescue these deﬁcits in other C. elegans mutants defective in osmosensation and mechanosensation (including OCR-2, bespeaking the speciﬁcity of the observed response). A point mutation in the pore loop of TRPV4, M680K, virtually eliminated rescue, indicating that TRPV4 functions as an ion channel. In an attempt to recapitulate the properties of the mammalian channel in the nociceptive behavior of the worm, it was found that the sensitivity for osmotic stimuli and the effect of temperature on the avoidance responses of osm9 ash::trpv4 worms more closely resembled properties of mammalian TRPV4 than that of normal worms. These data suggest that TRPV4 functions as an osmotically and mechanically gated channel, and that, in this model, TRPV4 directs the osmotic and mechanical avoidance behavior of the worm. TRPV4 does not rescue the odorant avoidance deﬁcit of osm-9 mutant worms, where G-protein-coupled receptors function as odorant sensors, and TRPV4 did not function downstream of other known mutations that affect nose touch and osmotic avoidance. In aggregate, these ﬁndings and considerations suggest that mammalian TRPV4 functioned as a component of the osmotic and mechanical sensor. TRPV4 was de facto expressed only in ASH, a single sensory neuron, where the mammalian protein, with a similarity to OSM-9 of approximately 25 percent, was trafﬁcked correctly to the ASH sensory cilia, a distance of about 50–100 micrometers! The rescue was speciﬁc (not for OCR-2, not by mammalian TRPV1), and it respected genetically deﬁned boundaries for osmotic and nose-touch avoidance. On the other hand, this study leads to stimulating questions. Whereas TRPV4 restores responsiveness to hyperosmotic stimuli in C. elegans osm-9 mutants, it is only gated by hypoosmotic stimuli in transfected mammalian cells. The reasons for this are not understood. One possibility is suggested by the results of a study where a mechanosensitive ion channel, gramicidin A, behaved either as a stretch-inactivated or as a stretch-activated channel depending on the lipid composition of the surrounding lipid bilayer [47]. An alternate possibility is that TRPV4 forms heteromultimeric complexes with other proteins, as was recently shown for the MEC proteins, and that this multiplex has different properties [15,29]. TRP ion channels are known to form heteromeric complexes with related family members [78,79]. OCR-2 and OSM-9 are the only C. elegans TRPV family members

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FIGURE 22.1 (From reference 45) Schematic representations illustrating how signal transduction in sensory (nerve) cells in response to odorant (A), osmotic (B) and mechanical (C) stimuli could possibly function. (A) The odorant activates the TRPV ion channel via a G-protein-coupled receptor mechanism. Such a mechanism is operative in the ASH sensory neuron of C.elegans in response to, e.g., 8-octanone, a repulsive odorant. Intracellular signaling cascades down-stream of the G-protein-coupled receptor activate the TRPV channel, OSM-9 or OCR-2. Calcium inﬂux through the TRPV channel serves as an ampliﬁcation mechanism, which is required for this signaling pathway to lead to the stereotypical withdrawal response. (B) This drawing represents two possibilities how tonicity signaling could work. In one alternative scenario, depicted on the right-hand side, the TRPV channel functions down-stream of a–yet unknown–osmotic stimulus transduction mechanism, which is directly activated by a change in tonicity. This is conceptually related to what is depicted in (A). Intracellular signaling via phosphorylation (de-phosphorylation)-dependent pathways activates the TRPV channel. For heterologous cellular expression systems, two groups have obtained data, contradictory in its detail, that suggest phosphorylation of TRPV4 to be of relevance [72,78]. On the left-hand side of the representation, note another scenario where the TRPV channel is at the top of the signaling cascade, i.e. it is directly activated by a change in tonicity, which in turn leads to an altered mechanical tension of the cytoplasma membrane via volume change.

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that are expressed in ASH neurons, and OCR-2 expression is essential for TRPV4 to rescue the sensory defects of osm-9 worms [47,66,67]. Related to this study, it was recently reported that TRPV2 could rescue one particular deﬁcit of the ocr-2 mutant, namely the dramatic downregulation of serotonin biosynthesis in the sensory ADF neuron, but mammalian TRPV2, unlike TRPV4 directing behavior in osm-9, did not complement the osmotic avoidance reaction that was lacking in ocr-2 mutants [61,80]. Common to these investigations is the conservation of TRPV signaling across phyla that were separated by several hundred million years of molecular evolution, and this in view of low sequence homology! In reference to the Drosophila TRPV channels, NAN and IAV, the reader is directed to original papers [28,38] and reviews [45,70]. One interesting generalization appears to be that a null mutant in an invertebrate TRPV channel has a stronger phenotype than a mutation in a mammalian TRPV channel.

OUTLOOK FOR FUTURE RESEARCH ON TRP CHANNELS Apart from the unexpected turns (the inevitability of those to occur renders the blossoming ﬁeld of TRP channels even more appealing than it already is [53,54]), the obvious topic for the (near) future is the investigation of the functional significance of protein–protein interactions of TRP(V) ion channels with known and tobe-discovered interaction partners. For example, a very interesting example of protein–protein interactions of TRPV4 splice variants from airway epithelia was reported [5], highlighting the need to also study the complexity of trp genes and their variants. In addition, there is the obvious potential for TRP channels as targets for therapy [30,39].

ACKNOWLEDGMENT The author was supported by a K08 Career Development Award of the National Institutes of Mental Health, by funding from the Whitehall Foundation (Palm Springs, FL), American Federation for Aging Research (New York, NY), the Klingenstein Fund (New York, NY), and Duke University (Durham, NC). FIGURE 22.1 (Continued) Note that the two alternatives need not be mutually exclusive. Apart from phosphorylation of the TRPV channel, which could possibly be of relevance in vivo, a direct physical linkage of the TRPV channel to the cytoskeleton, to the extracellular matrix and to the lipids of the plasma membrane adjacent to the channel has to be entertained. (C) This drawing represents two possibilities how mechanotransduction could work. Here, depicted on the right-hand side, an unknown mechanotransduction channel responds directly to the mechanical stimulus with calcium inﬂux. This activity and the subsequent signal transduction are modulated more indirectly by the TRPV channel, which acts on the unknown transduction channel, onto the biophysical properties of the membrane, and via other, yetunknown intracellular signaling mechanisms. The left-hand side depicts another alternative. Here, the TRPV channel is the mechanotransducer itself (i.e., it is activated directly via mechanical stimulation).

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REFERENCES [1]

[2]

[3]

[4]

[5]

[6]

[7] [8] [9]

[10] [11]

[12] [13]

[14]

[15] [16]

[17]

N. Alessandri-Haber, O.A. Dina, J.J. Yeh, C.A. Parada, D.B. Reichling, and J.D. Levine, Transient receptor potential vanilloid 4 is essential in chemotherapy-induced neuropathic pain in the rat, J. Neurosci. 24 (2004) 4444–4452. N. Alessandri-Haber, E. Joseph, O.A. Dina, W. Liedtke, and J.D. Levine, TRPV4 mediates pain-related behavior induced by mild hypertonic stimuli in the presence of inﬂammatory mediator, Pain 118 (2005) 70–79. Y.N. Andrade, J. Fernandes, E. Vazquez, J.M. Fernandez-Fernandez, M. Arniges, T.M. Sanchez, M. Villalon, and M.A. Valverde, TRPV4 channel is involved in the coupling of ﬂuid viscosity changes to epithelial ciliary activity, J. Cell Biol. 168 (2005) 869–874. A. Apostolidis, C.M. Brady, Y. Yiangou, J. Davis, C.J. Fowler, and P. Anand, Capsaicin receptor TRPV1 in urothelium of neurogenic human bladders and effect of intravesical resiniferatoxin, Urology 65 (2005) 400–405. M. Arniges, J.M. Fernandez-Fernandez, N. Albrecht, M. Schaefer, and M.A. Valverde, Human TRPV4 channel splice variants revealed a key role of ankyrin domains in multimerization and trafﬁcking, J. Biol. Chem. (2005). M. Arniges, E. Vazquez, J.M. Fernandez-Fernandez, and M.A. Valverde, Swellingactivated Ca2+ entry via TRPV4 channel is defective in cystic ﬁbrosis airway epithelia, J. Biol. Chem. 279 (2004) 54062–54068. C.I. Bargmann and J.M. Kaplan, Signal transduction in the Caenorhabditis elegans nervous system, Annu. Rev. Neurosci. 21 (1998) 279–308. D. Becker, C. Blase, J. Bereiter-Hahn, and M. Jendrach, TRPV4 exhibits a functional role in cell-volume regulation, J. Cell Sci. 118 (2005) 2435–2440. L.A. Birder, Y. Nakamura, S. Kiss, M.L. Nealen, S. Barrick, A.J. Kanai, E. Wang, G. Ruiz, W.C. De Groat, G. Apodaca, S. Watkins, and M.J. Caterina, Altered urinary bladder function in mice lacking the vanilloid receptor TRPV1, Nat. Neurosci. 5 (2002) 856–860. M.J. Caterina and D. Julius, Sense and speciﬁcity: a molecular identity for nociceptors, Curr. Opin. Neurobiol. 9 (1999) 525–530. M.J. Caterina, A. Lefﬂer, A.B. Malmberg, W.J. Martin, J. Trafton, K.R. PetersenZeitz, M. Koltzenburg, A.I. Basbaum, and D. Julius, Impaired nociception and pain sensation in mice lacking the capsaicin receptor, Science 288 (2000) 306–313. M.J. Caterina and C. Montell, Take a TRP to beat the heat, Genes Dev. 19 (2005) 415–418. M.J. Caterina, T.A. Rosen, M. Tominaga, A.J. Brake, and D. Julius, A capsaicinreceptor homologue with a high threshold for noxious heat, Nature 398 (1999) 436–441. M.J. Caterina, M.A. Schumacher, M. Tominaga, T.A. Rosen, J.D. Levine, and D. Julius, The capsaicin receptor: a heat-activated ion channel in the pain pathway, Nature 389 (1997) 816–824. M. Chalﬁe, Touch receptor development and function in Caenorhabditis elegans, J. Neurobiol. 24 (1993) 1433–1441. C.L. Chan, P. Facer, J.B. Davis, G.D. Smith, J. Egerton, C. Bountra, N.S. Williams, and P. Anand, Sensory ﬁbres expressing capsaicin receptor TRPV1 in patients with rectal hypersensitivity and faecal urgency, Lancet 361 (2003) 385–391. Q. Chang, S. Hoefs, A.W. van der Kemp, C.N. Topala, R.J. Bindels, and J.G. Hoenderop, The beta-glucuronidase klotho hydrolyzes and activates the TRPV5 channel, Science 310 (2005) 490–493.

TRPV Channels’ Function in Osmo- and Mechanotransduction

315

[18] D.E. Clapham, TRP channels as cellular sensors, Nature 426 (2003) 517–524. [19] H.A. Colbert, T.L. Smith, and C.I. Bargmann, OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans, J. Neurosci. 17 (1997) 8259–8269. [20] D.P. Corey, New TRP channels in hearing and mechanosensation, Neuron 39 (2003) 585–588. [21] D.J. Cosens and A. Manning, Abnormal electroretinogram from a Drosophila mutant, Nature 224 (1969) 285–287. [22] A.J. Culshaw, S. Bevan, M. Christiansen, P. Copp, A. Davis, C. Davis, A. Dyson, E.K. Dziadulewicz, L. Edwards, H. Eggelte, A. Fox, C. Gentry, A. Groarke, A. Hallett, T.W. Hart, G.A. Hughes, S. Knights, P. Kotsonis, W. Lee, I. Lyothier, A. McBryde, P. McIntyre, G. Paloumbis, M. Panesar, S. Patel, M.P. Seiler, M. Yaqoob, and K. Zimmermann, Identiﬁcation and biological characterization of 6-aryl-7-isopropylquinazolinones as novel TRPV1 antagonists that are effective in models of chronic pain, J. Med. Chem. 49 (2006) 471–474. [23] J.B. Davis, J. Gray, M.J. Gunthorpe, J.P. Hatcher, P.T. Davey, P. Overend, M.H. Harries, J. Latcham, C. Clapham, K. Atkinson, S.A. Hughes, K. Rance, E. Grau, A.J. Harper, P.L. Pugh, D.C. Rogers, S. Bingham, A. Randall, and S.A. Sheardown, Vanilloid receptor-1 is essential for inﬂammatory thermal hyperalgesia, Nature 405 (2000) 183–187. [24] O.A. Dina, X. Chen, D. Reichling, and J.D. Levine, Role of protein kinase C epsilon and protein kinase A in a model of paclitaxel-induced painful peripheral neuropathy in the rat, Neuroscience 108 (2001) 507–515. [25] O.A. Dina, C.A. Parada, J. Yeh, X. Chen, G.C. McCarter, and J.D. Levine, Integrin signaling in inﬂammatory and neuropathic pain in the rat, Eur. J. Neurosci. 19 (2004) 634–642. [26] P. Dinis, A. Charrua, A. Avelino, M. Yaqoob, S. Bevan, I. Nagy, and F. Cruz, Anandamide-evoked activation of vanilloid receptor 1 contributes to the development of bladder hyperreﬂexia and nociceptive transmission to spinal dorsal horn neurons in cystitis, J. Neurosci. 24 (2004) 11253–11263. [27] X. Gao, L. Wu, and R.G. O'Neil, Temperature-modulated diversity of TRPV4 channel gating: activation by physical stresses and phorbol ester derivatives through protein kinase C–dependent and –independent pathways, J. Biol. Chem. 278 (2003) 27129–27137. [28] Z. Gong, W. Son, Y.D. Chung, J. Kim, D.W. Shin, C.A. McClung, Y. Lee, H.W. Lee, D.J. Chang, B.K. Kaang, H. Cho, U. Oh, J. Hirsh, M.J. Kernan, and C. Kim, Two interdependent TRPV channel subunits, inactive and Nanchung, mediate hearing in Drosophila, J. Neurosci. 24 (2004) 9059–9066. [29] M.B. Goodman, G.G. Ernstrom, D.S. Chelur, R. O'Hagan, C.A. Yao, and M. Chalﬁe, MEC-2 regulates C. elegans DEG/ENaC channels needed for mechanosensation, Nature 415 (2002) 1039–1042. [30] T. Gudermann and V. Flockerzi, TRP channels as new pharmacological targets, Naunyn Schmiedebergs Arch. Pharmacol. 371 (2005) 241–244. [31] A.D. Guler, H. Lee, T. Iida, I. Shimizu, M. Tominaga, and M. Caterina, Heat-evoked activation of the ion channel, TRPV4, J. Neurosci. 22 (2002) 6408–6414. [32] M.J. Gunthorpe, C.D. Benham, A. Randall, and J.B. Davis, The diversity in the vanilloid (TRPV) receptor family of ion channels, Trends Pharmacol. Sci. 23 (2002) 183–191. [33] A.J. Hudspeth, How the ear's works work, Nature 341 (1989) 397–404. [34] A.J. Hudspeth, How the ear's works work: mechanoelectrical transduction and ampliﬁcation by hair cells, C. R. Biol. 328 (2005) 155–162.

316 [35] [36]

[37] [38]

[39]

[40]

[41]

[42]

[43]

[44] [45]

[46]

[47]

[48] [49]

[50] [51] [52]

TRP Ion Channel Function in Sensory Transduction A.J. Hudspeth and P.G. Gillespie, Pulling springs to tune transduction: adaptation by hair cells, Neuron 12 (1994) 1–9. M. Kanzaki, Y.Q. Zhang, H. Mashima, L. Li, H. Shibata, and I. Kojima, Translocation of a calcium-permeable cation channel induced by insulin-like growth factor-I, Nat. Cell Biol. 1 (1999) 165–170. J.M. Kaplan and H.R. Horvitz, A dual mechanosensory and chemosensory neuron in Caenorhabditis elegans, Proc. Natl. Acad. Sci. USA 90 (1993) 2227–2231. J. Kim, Y.D. Chung, D.Y. Park, S. Choi, D.W. Shin, H. Soh, H.W. Lee, W. Son, J. Yim, C.S. Park, M.J. Kernan, and C. Kim, A TRPV family ion channel required for hearing in Drosophila, Nature 424 (2003) 81–84. J.E. Krause, B.L. Chenard, and D.N. Cortright, Transient receptor potential ion channels as targets for the discovery of pain therapeutics, Curr. Opin. Invest. Drugs 6 (2005) 48–57. M. Lazzeri, M.G. Vannucchi, C. Zardo, M. Spinelli, P. Beneforti, D. Turini, and M.S. Faussone-Pellegrini, Immunohistochemical evidence of vanilloid receptor 1 in normal human urinary bladder, Eur. Urol. 46 (2004) 792–798. H. Lee, T. Iida, A. Mizuno, M. Suzuki, and M.J. Caterina, Altered thermal selection behavior in mice lacking transient receptor potential vanilloid 4, J. Neurosci. 25 (2005) 1304–1310. X. Liu, B.B. Bandyopadhyay, T. Nakamoto, B.B. Singh, W. Liedtke, J.E. Melvin, and I.S. Ambudkar, A role for AQP5 in activation of TRPV4 by hypotonicity: concerted invovement of AQP5 and TRPV4 in regulation of volume recovery, J. Biol. Chem. 281 (2006) 15485–15495. W. Liedtke, Y. Choe, M.A. Marti-Renom, A.M. Bell, C.S. Denis, A. Sali, A.J. Hudspeth, J.M. Friedman, and S. Heller, Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor, Cell 103 (2000) 525–535. W. Liedtke and J.M. Friedman, Abnormal osmotic regulation in trpv4−/− mice, Proc. Natl. Acad. Sci. USA 100 (2003) 13698–13703. W. Liedtke and C. Kim, Functionality of the TRPV subfamily of TRP ion channels: add mechano-TRP and osmo-TRP to the lexicon! Cell Mol. Life Sci. 62 (2005) 2985–3001. W. Liedtke, D.M. Tobin, C.I. Bargmann, and J.M. Friedman, Mammalian TRPV4 (VR-OAC) directs behavioral responses to osmotic and mechanical stimuli in Caenorhabditis elegans, Proc. Natl. Acad. Sci. USA 100 Suppl. 2 (2003) 14531–14536. B. Martinac and O.P. Hamill, Gramicidin A channels switch between stretch activation and stretch inactivation depending on bilayer thickness, Proc. Natl. Acad. Sci. USA 99 (2002) 4308–4312. A. Mizuno, N. Matsumoto, M. Imai, and M. Suzuki, Impaired osmotic sensation in mice lacking TRPV4, Am. J. Physiol. Cell Physiol. 285 (2003) C96–101. K. Muraki, Y. Iwata, Y. Katanosaka, T. Ito, S. Ohya, M. Shigekawa, and Y. Imaizumi, TRPV2 is a component of osmotically sensitive cation channels in murine aortic myocytes, Circ. Res. 93 (2003) 829–838. H. Mutai and S. Heller, Vertebrate and invertebrate TRPV-like mechanoreceptors, Cell Calcium 33 (2003) 471–478. R.S. Naeini, M.F. Witty, P. Seguela, and C.W. Bourque, An N-terminal variant of Trpv1 channel is required for osmosensory transduction, Nat. Neurosci. (2005). K. Nagata, A. Duggan, G. Kumar, and J. García-Añoveros, Nociceptor and hair cell transducer properties of TRPA1, a channel for pain and hearing, J. Neurosci. 25 (2005) 4052–4061.

TRPV Channels’ Function in Osmo- and Mechanotransduction [53] [54] [55] [56]

[57]

[58]

[59]

[60]

[61]

[62]

[63]

[64] [65]

[66]

[67]

317

B. Nilius and S.O. Sage, TRP channels: novel gating properties and physiological functions, J. Physiol. 567 (2005) 33–34. B. Nilius, T. Voets, and J. Peters, TRP channels in disease, Sci. STKE 2005 (2005) re8. A. Patapoutian, TRP channels and thermosensation, Chem. Senses 30 Suppl. 1 (2005) i193–i194. A.M. Peier, A.J. Reeve, D.A. Andersson, A. Moqrich, T.J. Earley, A.C. Hergarden, G.M. Story, S. Colley, J.B. Hogenesch, P. McIntyre, S. Bevan, and A. Patapoutian, A heat-sensitive TRP channel expressed in keratinocytes, Science 296 (2002) 2046–2049. J.D. Pomonis, J.E. Harrison, L. Mark, D.R. Bristol, K.J. Valenzano, and K. Walker, N-(4-Tertiarybutylphenyl)-4-(3-cholorphyridin-2-yl)tetrahydropyrazine-1(2H)-carboxamide (BCTC), a novel, orally effective vanilloid receptor 1 antagonist with analgesic properties: II. in vivo characterization in rat models of inﬂammatory and neuropathic pain, J. Pharmacol. Exp. Ther. 306 (2003) 387–393. W. Rong, K. Hillsley, J.B. Davis, G. Hicks, W.J. Winchester, and D. Grundy, Jejunal afferent nerve sensitivity in wild-type and TRPV1 knockout mice, J. Physiol. 560 (2004) 867–881. R.S. Scotland, S. Chauhan, C. Davis, C. De Felipe, S. Hunt, J. Kabir, P. Kotsonis, U. Oh, and A. Ahluwalia, Vanilloid receptor TRPV1, sensory C-ﬁbers, and vascular autoregulation: a novel mechanism involved in myogenic constriction, Circ. Res. 95 (2004) 1027–1034. G.D. Smith, M.J. Gunthorpe, R.E. Kelsell, P.D. Hayes, P. Reilly, P. Facer, J.E. Wright, J.C. Jerman, J.P. Walhin, L. Ooi, J. Egerton, K.J. Charles, D. Smart, A.D. Randall, P. Anand, and J.B. Davis, TRPV3 is a temperature-sensitive vanilloid receptor-like protein, Nature 418 (2002) 186–190. I. Sokolchik, T. Tanabe, P.F. Baldi, and J.Y. Sze, Polymodal sensory function of the Caenorhabditis elegans OCR-2 channel arises from distinct intrinsic determinants within the protein and is selectively conserved in mammalian TRPV proteins, J. Neurosci. 25 (2005) 1015–1023. R.J. Stein, S. Santos, J. Nagatomi, Y. Hayashi, B.S. Minnery, M. Xavier, A.S. Patel, J.B. Nelson, W.J. Futrell, N. Yoshimura, M.B. Chancellor, and F. De Miguel, Cool (TRPM8) and hot (TRPV1) receptors in the bladder and male genital tract, J. Urol. 172 (2004) 1175–1178. R. Strotmann, C. Harteneck, K. Nunnenmacher, G. Schultz, and T.D. Plant, OTRPC4, a nonselective cation channel that confers sensitivity to extracellular osmolarity, Nat. Cell Biol. 2 (2000) 695–702. M. Suzuki, A. Mizuno, K. Kodaira, and M. Imai, Impaired pressure sensation in mice lacking TRPV4, J. Biol. Chem. 278 (2003) 22664–22668. W. Tian, M. Salanova, H. Xu, J.N. Lindsley, T.T. Oyama, S. Anderson, S. Bachmann, and D.M. Cohen, Renal expression of osmotically responsive cation channel TRPV4 is restricted to water-impermeant nephron segments, Am. J. Physiol. Renal Physiol. 287 (2004) F17–24. D. Tobin, D.M. Madsen, A. Kahn-Kirby, E. Peckol, G. Moulder, R. Barstead, A.V. Maricq, and C.I. Bargmann, Combinatorial expression of TRPV channel proteins deﬁnes their sensory functions and subcellular localization in C. elegans neurons, Neuron 35 (2002) 307–318. D.M. Tobin and C.I. Bargmann, Invertebrate nociception: behaviors, neurons and molecules, J. Neurobiol. 61 (2004) 161–174.

318 [68]

[69] [70]

[71]

[72] [73]

[74] [75]

[76]

[77]

[78]

[79] [80]

[81] [82]

TRP Ion Channel Function in Sensory Transduction H. Todaka, J. Taniguchi, J. Satoh, A. Mizuno, and M. Suzuki, Warm temperaturesensitive transient receptor potential vanilloid 4 (TRPV4) plays an essential role in thermal hyperalgesia, J. Biol. Chem. 279 (2004) 35133–35138. M. Tominaga and M.J. Caterina, Thermosensation and pain, J. Neurobiol. 61 (2004) 3–12. J. Vriens, G. Owsianik, T. Voets, G. Droogmans, and B. Nilius, Invertebrate TRP proteins as functional models for mammalian channels, Pﬂügers Arch. 449 (2004) 213–226. J. Vriens, H. Watanabe, A. Janssens, G. Droogmans, T. Voets, and B. Nilius, Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4, Proc. Natl. Acad. Sci. USA (2003). R.G. Walker, A.T. Willingham, and C.S. Zuker, A Drosophila mechanosensory transduction channel, Science 287 (2000) 2229–2234. H. Watanabe, J. Vriens, S.H. Suh, C.D. Benham, G. Droogmans, and B. Nilius, Heatevoked activation of TRPV4 channels in a HEK293 cell expression system and in native mouse aorta endothelial cells, J. Biol. Chem. 277 (2002) 47044–47051. U. Wissenbach, M. Bodding, M. Freichel, and V. Flockerzi, Trp12, a novel Trp-related protein from kidney, FEBS Lett. 485 (2000) 127–134. H. Xu, Y. Fu, W. Tian, and D.M. Cohen, Glycosylation of the osmoresponsive transient receptor potential channel TRPV4 on Asn-651 inﬂuences membrane trafﬁcking, Am. J. Physiol. Renal Physiol. (2005). H. Xu, I.S. Ramsey, S.A. Kotecha, M.M. Moran, J.A. Chong, D. Lawson, P. Ge, J. Lilly, I. Silos-Santiago, Y. Xie, P.S. DiStefano, R. Curtis, and D.E. Clapham, TRPV3 is a calcium-permeable temperature-sensitive cation channel, Nature 418 (2002) 181–186. H. Xu, H. Zhao, W. Tian, K. Yoshida, J.B. Roullet, and D.M. Cohen, Regulation of a transient receptor potential (TRP) channel by tyrosine phosphorylation. SRC family kinase-dependent tyrosine phosphorylation of TRPV4 on TYR-253 mediates its response to hypotonic stress, J. Biol. Chem. 278 (2003) 11520–11527. X.Z. Xu, F. Chien, A. Butler, L. Salkoff, and C. Montell, TRPgamma, a drosophila TRP-related subunit, forms a regulated cation channel with TRPL, Neuron 26 (2000) 647–657. X.Z. Xu, H.S. Li, W.B. Guggino, and C. Montell, Coassembly of TRP and TRPL produces a distinct store-operated conductance, Cell 89 (1997) 1155–1164. S. Zhang, I. Sokolchik, G. Blanco, and J.Y. Sze, Caenorhabditis elegans TRPV ion channel regulates 5HT biosynthesis in chemosensory neurons, Development 131 (2004) 1629–1638. S. Ciura and C.W. Bourque, Impaired detection of hyperosmotic stimuli in the OVLT of TRPV1 KO mice, The Physiologist 48 (2005) 418. V.K. Sidhaye, A.D. Güler, K.S. Schweitzer, F. D’Alessio, M.J. Caterina, and L.S. King, Transient receptor potential vanilloid 4 regulates aquaporin-5 abundance under hypotonic conditions, Proc. Natl. Acad. Sci. USA 103 (2006) 4747–4752.

23

TRP Channel Trafﬁcking Rosa Planells-Cases Centro de Investigación Príncipe Felipe

Antonio Ferrer-Montiel Universidad Miguel Hernández

CONTENTS TRP Deﬁnitions and Families ...............................................................................322 TRP Translocation as an Activation and Regulatory Mechanism ........................324 Activation of SOC Channels by Protein Translocation to the Plasma Membrane...........................................................................324 Functional Modulation of TRPL and TRP3 Channels .............................325 Agonist-Induced Translocation of TRPC Channels..................................325 Modulation of TRPV Channel Activity ....................................................326 Concluding Remarks .............................................................................................327 Acknowledgments..................................................................................................328 References..............................................................................................................329 Cellular function, in particular neuronal activity, is the result of an orchestrated interplay between membrane receptors at the cell surface and intracellular signaling proteins. Membrane receptors transduce external signals into cellular responses by activating cytosolic metabolic pathways. Regulation of the spatial and temporal distribution of membrane receptors begins to be recognized as an important mechanism for controlling the magnitude and time course of cellular signaling.1 For instance, long-term changes in neuronal plasticity are the result of transcriptional and translational regulation of protein biogenesis and trafﬁcking. Similarly, shortterm changes in synaptic function can be achieved by altering the levels of key surface proteins involved in synaptic transmission, such as neurotransmitter receptors and transporters.1 Thus, modulation of receptor function by trafﬁcking and redistribution plays a central role in synaptic plasticity.1–3 These processes increase the complexity of the signaling mechanisms in peripheral and central synapses. Protein trafﬁcking studies, carried out in yeast and mammalian cells, have revealed the underlying mechanism by which proteins travel between intracellular compartments and the plasma membrane. Membrane proteins are synthesized in the endoplasmic reticulum (ER), and ensuing posttranslational modiﬁcations such as glycosylation take place in both the ER and the Golgi apparatus. Appropriately 319

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folded and assembled membrane proteins are subsequently sorted out into vesicles for trafﬁcking and delivery to the cell surface by a constitutive or regulated pathway of vesicle exocytosis.4 The constitutive exocytotic route is primarily used by cells for turnover of both membrane proteins and lipids, thus ensuring the removal of aged or damaged membrane components. In contrast, the regulated secretory pathway is used by specialized cells, such as endocrine cells, neurons, acinar cells of the pancreas, mast cells, insulin-sensitive cells, and many other cell types, as a mechanism of cellular communication. Regulated exocytosis is produced only in response to a stimulus that provokes the mobilization, docking, and fusion of an intracellular pool of vesicles stored near the membrane region where they will fuse and release their components.1–5 Mechanistically, both types of exocytosis use a cascade of protein–protein interactions to ensure the efﬁcient delivery of their vesicle cargo to the membrane.2,5 At a molecular level, vesicle trafﬁc uses the SNAREs (soluble N-ethylmaleimidesensitive factor attachment protein receptors), a large family of proteins that are present on all organelles and mediate intracellular vesicle trafﬁcking and secretion (Figure 23.1).2,5 The interaction of complementary SNARE proteins found on opposing membranes exempliﬁes an attractive lock-and-key mechanism, which presumably underlies the remarkable speciﬁcity of vesicle docking to target membranes Synapse

Golgi

Microtubule

Endosome

TRP v-SNARE

t-SNARE

FIGURE 23.1 SNARE-dependent exocytosis of transport and recycling vesicles at a synapse. The two pathways of generating synaptic vesicles containing TRP channels are depicted: a pathway where vesicles are transported from the Golgi to the synaptic terminal, and the parallel pathway where vesicles are generated following the endocytotic pathway via an endosome.1–5 Synaptic vesicles are secreted by SNARE-dependent exocytosis (inset). The inset shows the steps of vesicle docking and fusion. The v- and t-SNAREs assemble into a four-helix bundle, promoting fusion of the vesicle to the plasma membrane and thus incorporation of the receptor to the surface membrane. Once fused, SNAREs and the TRP channels are recycled.

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321

and their consequent fusion.5 For instance, the interaction of the vesicle membrane SNARE (v-SNARE) synaptobrevin 2, and the corresponding SNAREs in the target membrane (t-SNAREs), syntaxin 1 and SNAP25, results in the formation of a remarkably stable ternary complex that ensures the docking and fusion of vesicles at the active zone upon receiving the stimulus, normally an increase in the intracellular concentration of Ca2+ ([Ca2+]i).5 Similarly, molecular interactions of synaptobrevin 2 analogues with syntaxin 1 and SNAP25-like target membrane proteins are thought to be essential for vesicle targeting and fusion in all eukaryotes.2 It is quite well established that a signiﬁcant increase in [Ca2+]i is required for triggering the SNARE-mediated fusion of secretory vesicles, although SNARE-dependent exocytosis of vesicles, in particular recycling vesicles, may occur at resting [Ca2+]i by the activation of other signaling pathways.2 Nonetheless, the rate of vesicle recycling is regulated by changes in the [Ca2+]i. Vesicle exocytosis is a highly regulated process counterbalanced by endocytosis of membrane components that ensures constant plasma membrane content. Internalization of receptors and transporters is mediated by a general clathrindependent pathway that moves endocytosed proteins to the endosome for appropriate sorting into vesicles, either for release or for transport depending on their ﬁnal destination.3,4 The endosome also provides a mechanism to maintain the

S4

S2 S1

S3

S6

S5

TRP domain

CaM

PIP2

PDZ

CC Ankyrin domains

PLIK kinase

Ca2+ K+ Na+

FIGURE 23.2 Molecular model of the TRP receptor subunit. The ﬁgure displays a membrane domain composed of six transmembrane segments (S1–S6) with an amphipathic region between the ﬁfth and sixth segment that forms the channel conductive pore. The protein also has cytoplasmic N- and C-termini. In the N-terminus, TRP channels may exhibit ankyrin domains and a coiled-coil domain.11,12 These proteins display a cytosolic C-terminus domain that, depending on the TRP subfamily, may display phosphoinositide (PIP2), calmodulinbinding (CAM), PLIK kinase, and PDZ domains.11,12 In addition, the C-end may also exhibit a TRP domain that may contribute to the association of receptor subunits.16 The higher Ca2+ permeability with respect to Na+ and K+ is also depicted.

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proper composition of release vesicle proteins by adding newly synthesized vesicle proteins, removing aged polypeptides for degradation, and separating nonsecretory vesicle proteins.3 Recent evidence suggests that synaptic vesicles are most likely formed at the nerve terminal by endocytosis. This novel notion considers that newly synthesized membrane proteins trafﬁc to the nerve terminal in transport packets and become incorporated into synaptic vesicles at the synapse, thus indicating that synaptic vesicle formation relies primarily on an endocytotic pathway.3,4 This hypothesis is particularly attractive for the regulation of the surface expression of ion channels, neurotransmitter receptors, and transporters presynaptically due to the lack of biosynthetic pools of membrane proteins available in the synapse.4 Traditionally, the constitutive exocytotic pathway has been associated to the trafﬁc of membrane proteins to the cell surface, while the regulated secretory pathway has been implicated in the activity-dependent release of neurotransmitters, neuropeptides, and hormones. However, in the past decade emerged the notion that the surface expression of membrane proteins, especially ion channels, receptors, and transporters, may be mediated by a regulated release pathway as well.2–4 Indeed, cumulative evidence indicates that the function of these plasma membrane proteins can be increased or decreased by accommodating the level of the surface-expressed protein. Indeed, dynamic regulation of the rate of either insertion or retrieval or both of integral membrane proteins in response to stimuli embodies the strategic regulation of their surface expression.3,4 Examples of receptors and transporters that are modulated by vesicle trafﬁcking are the glucose transporter (GluT4),6 Aquaporin 2 (AQP2),7 epithelial sodium channel (ENac),8 cystic ﬁbrosis transmembrane conductance regulator (CFTR),8 as well as ionotropic ligand-gated receptors such as -amino3-hydroxy-5-methylisoxazole-4-propionate (AMPA), N-methyl-D-aspartate (NMDA), gamma-aminobutyrate (GABAA), and the purinergic receptor P2X4.9 In addition, several studies have demonstrated that this signaling pathway is also commonly used by members of the transient receptor potential (TRP) family of ion channels for modulation of their physiological function.10

TRP DEFINITIONS AND FAMILIES The TRP channels are a superfamily of ion channels that play a wide diversity of physiological functions and are present in many tissues and almost all cell types.11,12 Most TRP channels are nonselective cation channels with low voltage dependence. TRP channels use a wide variety of activation and regulatory mechanisms and carry out functions as diverse as thermosensation, pheromone reception, magnesium homeostasis, and regulation of vascular tone. Interestingly, these channels are considered molecular gateways in sensory and regulatory systems. The founder member of the family, the TRP channel, was cloned from the ﬂy Drosophila. The phenotype of the ﬂy, trp, was caused by a sustained response in the electroretinogram in response to continuous light, instead of a transient response, with a concomitant decrease in the light-induced Ca2+ inﬂux. Mutation in the gene TRP caused the defect in a Ca2+ channel required for Ca2+ inﬂux and loss of the transient voltage response.13 A second gene, trp-like (trpl), with ~40 percent identity

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323

with trp, encodes a second class of light-sensitive channel that is responsible for the light-induced current.14 Since the cloning of the ﬂy TRP, more than 50 different gene products have been reported from fungal genomes to upper mammals. Within the animal kingdom, different TRP genes are well conserved. In mice and humans, 28 TRP homologue genes have been described, whereas in invertebrates there are close to 15 members. Based on their amino acid sequence homology, mammalian TRP channels are divided into six subfamilies, namely the “canonical” TRPC (composed of TRPC1 to TRPC7), “vanilloid” TRPV (with TRPV1 to TRPV6), “melastatin-related” TRPM (with TRPM1 to TRPM8), “polycystin-related” TRPP, “mucolipin-related” TRPML and “ankyrin” TRPA.11,12,15 An additional family, TRPN, has been described in Drosophila.15 TRP proteins have a common topology of six transmembrane segments (S1–S6) with a pore region between the ﬁfth and sixth segment, and cytoplasmic N- and Ctermini, reminiscent of voltage-dependent K+ (Kv) channels (Figure 23.2). The charged residues in the putative S4 helix, which usually underlie voltage-dependent gating to Kv channels, are replaced by noncharged amino acids. The length of the cytoplasmic domains varies within subfamilies as well as the structural and functional domains associated to them. As for Kv channels, a functional unit of TRP channel requires tetrameric assembly of homomeric or heteromeric TRP polypeptides. TRPV and TRPC members contain two to four ankyrin domains at the Nterminus that are thought to interact with the cytoskeleton as well as with other cytosolic proteins.11 In the C-terminus domain, proximal to the S6 transmembrane segment, the TRP domain, a highly conserved segment of 25 amino acids (six of which are referred to as the TRP box), is thought to conform an association domain involved in subunit multimerization to give rise to an active channel.16 In addition, a putative calmodulin (CaM)-binding domain in the COOH terminal region has been reported that binds CaM in a Ca2+-dependent manner.17,18 Mammalian TRPCs are closest in sequence to Drosophila TRP and TRPL and were the ﬁrst to be cloned. Initially, these channels were proposed to function as store-operated Ca2+ channels (SOCs).19 SOC channels represent the primordial Ca2+ entry pathway. They activate upon Ca2+ release from internal stores either by the Ca2+ATPase inhibitor thapsigargin, or by the presence of ionomycin in Ca2+-free medium.19,20 Store depletion causes gating of plasma membrane Ca2+-permeable channels, resulting in Ca2+ inﬂux. SOCs are believed to replenish and maintain ER Ca2+ levels and to generate prolonged Ca2+ signals. The most intensely studied SOC is the Ca2+ release–activated Ca2+ current (ICRAC). ICRAC is a nonvoltage-activated and Ca2+ selective channel, inwardly rectifying at negative voltages, ﬁrst described in mast cells.20 ICRAC is not the only SOC channel because other store-operated inﬂux have been described with different biophysical properties in other tissues such as epidermal, endothelial, vascular smooth muscle, and neuronal cells. Although the molecular identity of SOC channels is yet unknown, it has been proposed that they are TRP proteins. Speciﬁcally, several TRPC channels have been shown to mediate Ca2+ entry with functional and biophysical properties that mirror those of SOC channels.21,22 In addition, the TRPV6 channel has a high Ca2+ selectivity that suggests a functional similarity to the CRAC channel, and in some studies activates in response to store

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TRP Ion Channel Function in Sensory Transduction

depletion,23,24 although this conclusion is still a matter of discussion.25,26 Thus, despite all these ﬁndings, the role of TRP channels in mediating Ca2+ entry in response to Ca2+ store depletion and receptor activation remains uncertain and controversial.

TRP TRANSLOCATION AS AN ACTIVATION AND REGULATORY MECHANISM TRP channels use a wide variety of activation and regulatory mechanisms to carry their functions. For instance, channel activity is affected by different physical stimuli such as light, temperature, osmolarity, mechanical force, or by different chemical or biochemical factors such as pH and external and internal ligands. These stimuli may directly activate the channels such as with TRPV1, which is gated by heat, acid, and vanilloids, or the stimuli may open the channels through a phospholipase C-inositol trisphosphate-mediated pathway.11,12 For instance, TRPC channels activate primarily in response to PLC-coupled receptors and interact with intracellular InsP3 receptors, suggesting that they can receive information directly from Ca2+ stores, although there is also evidence that TRPC channels can function independently of stores.11,12,15 Some members of the TRP channel family appear to be constitutively active channels because they are not gated by an identiﬁed stimulus. Cumulative evidence substantiates the tenet that regulated translocation of TRP proteins to the plasma membrane may be a potentially common, but poorly understood, mode of activation of these channels (Figure 23.1).10 This mode of activation has been particularly proposed for those TRP channels that are constitutively active, and whose overactivity may lead to Ca2+ overload, although the members of the TRP family that are regulated by this mechanism is augmenting. Regulated translocation of TRPcontaining vesicles could be envisioned as a strategy for brief, controlled increases in Ca2+ inﬂux. More generally, fast activity-dependent incorporation of TRP channels from a cytosolic vesicle pool to the plasma membrane, followed by endocytotic uptake, could exemplify an elegant mechanism of ion channel regulation.4 This process has been demonstrated for several members of the TRP channel family as described next.

ACTIVATION OF SOC CHANNELS PLASMA MEMBRANE

BY

PROTEIN TRANSLOCATION

TO THE

Although the molecular identity of SOC channels is yet elusive, there is evidence that TRP channels may underlie SOC-mediated Ca2+ inﬂux. These channels are activated by store depletion through a yet unknown mechanism. Three hypotheses have been proposed:19 (1) the existence of a diffusible factor released from the empty stores; (2) direct coupling of SOC channels in the plasma membrane with a protein sensor in the ER; and (3) vesicle-mediated translocation of SOC channels from a readily available, sub-plasmalemmal vesicle pool. Yao et al.,27 in a study of SOC activity in Xenopus oocytes, documented that SOC activity could be measured in the absence of a small messenger molecule. More interestingly, these authors showed that SOC function was affected by treatments that speciﬁcally abrogated vesicle trafﬁcking to and fusion with the plasma membrane. In particular, regulation of Rho A signiﬁcantly inﬂuenced the magnitude of the SOC-mediated ionic currents in

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325

oocytes.27 More important, SOC-dependent currents were inhibited 50 percent by treatment with botulinum neurotoxin A (BoNTA) and by a dominant-negative mutant of SNAP-25, thus indicating that SOC currents are due to translocation of channels to the cell surface by SNARE-mediated exocytosis. Similarly, BoNTA and tetanus neurotoxin (TeNT) reduced cyclopiazonic-induced, SOC-mediated Ca2+ entry into embryonic kidney cells,28 further substantiating the involvement of a regulated secretory pathway for SOC activation. However, the vesicular delivery hypothesis was questioned by Scott et al.,29 who reported that SOC-dependent Ca2+ ﬂuxes were insensitive to BoNTA, TeNT, and a dominant-negative mutant of NSF, suggesting that SOC channels and the machinery necessary to activate them are present in the plasma membrane prior to activation. Taken together, these results imply that trafﬁcking of SOC channels to the plasma membrane may contribute at least in part to channel gating, although further experimental evidence is needed to demonstrate this mechanism of activation.

FUNCTIONAL MODULATION

OF

TRPL

AND

TRP3 CHANNELS

Important evidence that a TRP channel undergoes dynamic translocation in a native system was obtained in ﬂy photoreceptor cells.30 In Drosophila, light regulates the subcellular distribution of TRPL channels between the rhabdomers and the plasma membrane, and it induces long-term adaptation to luminosity that modiﬁes the lightinduced, TRPL-mediated ionic currents. A pivotal ﬁnding was that the TRPL channel is translocated back and forth between the signaling membrane and the intracellular compartment by a light-regulated mechanism, resulting in a high-level rhabdomeral TRPL in the dark and a low level in the light.30 A physiological consequence of regulating the subcellular distribution of TRPL is that photoreceptors of ﬂies kept in the dark are sensitive to dim backgrounds, which allows photoreceptors enriched in TRPL to function better in darkness and faint background illumination. In Caenorhabditis elegans, the TRPC channel TRP-3, which is present in intracellular vesicles of spermatids, translocates to the plasma membrane during sperma activation, thus contributing to the acrosomal reaction.31 Akin to the TRPL, the regulated translocation of TRP-3 channels to the plasma membrane has profound physiological consequences. The molecular mechanisms underlying the regulated exocytosis of both TRPL and TRP-3 channels are yet elusive, although they may involve vesicle recycling and SNARE-dependent exocytosis.

AGONIST-INDUCED TRANSLOCATION

OF

TRPC CHANNELS

Three members of the TRPC subfamily of ion channels—TRPC3, TRPC5, and TRPC6—have been found to be regulated by an agonist-activated, SNARE-dependent vesicle translocation to the plasma membrane.32–34 In a nice study, Singh et al.32 found that VAMP-dependent, Ca2+-independent exocytosis regulates plasma membrane insertion of TRPC3 channels and contributes to agonist-stimulated Ca2+ inﬂux. These authors found that TRPC3-containing vesicles were localized immediately below the plasma membrane, and they demonstrated that agonist-induced TRPC3 translocation appears to be receptor speciﬁc because thapsigargin, a compound that releases Ca2+

326

TRP Ion Channel Function in Sensory Transduction

from the ER, did not change the surface expression of the channel. Protein translocation to the plasma membrane, but not channel trafﬁcking, was abrogated by cleavage of VAMP2 with TeNT, indicating a SNARE-dependent exocytosis.32 In contrast, although [Ca2+]i at or below resting modulated trafﬁcking of the TRPC3 vesicle to the plasma membrane, the agonist-induced surface expression of the protein was not produced by a rise in [Ca2+]i. In a similar study, Cayouette et al.33 found in a cell line stably expressing the TRPC6 channel that agonist-induced Ca2+ inﬂux was preceded by exocytotic insertion of TRPC6 proteins into the plasma membrane. Notably, they observed that externalization of TRPC6 channels occurred at an agonist concentration lower than that required for Ca2+ entry, implying that a signiﬁcant rise in [Ca2+]i is not implied. However, at variance with TRPC3, depletion of Ca2+ stores by thapsigargin caused the translocation of the channel to the cell surface, although it did not activate the channel.33 The molecular mechanism involved in the exocytosis of TRPC6 was not addressed in the study, although, akin to the TRPC3 channel, it may be mediated by SNARE proteins. Taken together, these ﬁndings provide pivotal information for understanding the molecular mechanism involved in agoniststimulated, G-protein-coupled Ca2+ inﬂux through these TRP channels. Both TRPC3 and TRPC6 studies provide compelling evidence of the existence of ligand-induced translocation of channels in the cell surface. These ﬁndings are further substantiated by the results reported for TRPC5. The work of Bezzerides et al.34 clearly demonstrated that stimulation of HEK293 cells expressing TRPC5 channels with EGF initiated the rapid incorporation of the channel from a plasmalemma pool of vesicles to the cell surface. This process was coined with the term rapid vesicular insertion of TRP (RiVIT).34 These authors were also able to uncover the molecular mechanism of channel translocation and found that the signaling pathway involved phosphatidylinositide 3-kinase, the Rho GTPase Rac1, and phosphatidylinositol-4 phosphate 5-kinase (PIP[5]K). Most remarkable, translocation of TRPC5 from the vesicular pool was observed in hippocampal neurons in culture. In this in vivo system, nerve growth factor (NGF), brain-derived growth factor (BDNF), and insulin growth factor (IGF) promoted the rapid exocytosis of TRPC5 into the neuronal membrane.34 Because TRPC5 interacts with VAMP and sinaptotagmin I in synaptic vesicles,35 it is reasonable to conclude that these proteins target the TRPC5containing vesicles to the plasma membrane, thus mediating the SNARE-dependent exocytotic insertion of the channel. Physiologically, the stimulus-induced increase of TRPC5 channels in the neuronal surface notably affected neurite extension,34,35 presumably by modulating the extent of Ca2+ inﬂux at the synaptic terminal. Thus, this elegant study demonstrates that regulated exteriorization of ion channels provides a mechanism for transient, local, self-limiting increases of Ca2+ through TRP channels. The insertion-retrieval of channels from the cell surface represents a clever strategy to avoid the consequences of Ca2+ overload that could occur through TRP channels that are constitutively active.

MODULATION

OF

TRPV CHANNEL ACTIVITY

TRPV channels have also been reported to be inserted into the plasma membrane in response to a stimulus. Indeed, the ﬁrst evidence suggesting exocytotic

TRP Channel Trafﬁcking

327

mechanism–mediated TRP channel activity was the demonstration that IGF induced the translocation of TRPV2 to the plasma membrane in cells heterologously overexpressing the channel.36,37 However, a study of hippocampal neurons in culture failed to observe IGF-induced translocation of TRPV2 to the plasma membrane.34 Thus, whether regulated exocytosis occurs in vivo is still under debate. The TRPV5 and TRPV6 channels are members of the TRPV channels that appear constitutively active and very selective for Ca2+. Therefore, akin to the TRPC family, it seems reasonable that the activity of these channels is also controlled by their regulated insertion into the plasma membrane when needed. Although no direct evidence for the secretion mechanism is available yet for TRPV5 and TRPV6 exocytosis, their interaction with the S100A-annexin 2 complex, which is associated with the cortical cytoskeleton, clearly suggests this sort of mechanism.38 Similar to other TRP channels, TRPV1 is arranged in major molecular complexes establishing high-order signaling networks that notably determine the response to external stimuli. These proteins’ networks are in turn the target of intracellular signaling pathways that modulate their composition, structure, and function. A yeast-two hybrid screen of a rat brain library using the N-terminus of TRPV1 as bait identiﬁed two synaptic vesicle proteins that were interacting partners of TRPV1: Snapin and Synaptotagmin IX.39 These proteins bind to the SNARE proteins and participate in neuronal exocytosis, suggesting that surface delivery of TRPV1 channels is a highly regulated, Ca2+-dependent exocytotic process (Figure 23.3). Indeed, the interaction of both vesicular proteins with TRPV1 appears temporal and seems uninvolved in the formation of the molecular complexes at the cell surface. However, these interactions are pivotal for the trafﬁcking and surface expression of TRPV1 channels in response to activation of intracellular pathways by inﬂammatory mediators.39,40 In support of this tenet, PKC-induced TRPV1 trafﬁcking to the plasma membrane was blocked by botulinum neurotoxins, indicating that PKC-induced sensitization of TRPV1 receptors is due at least in part to the regulated exocytosis of channels located in a reserve pool of cytosolic vesicles (Figure 23.3).39,40 A recent observation that botulinum neurotoxin attenuates heat hyperalgesia substantiates this hypothesis,41 although additional in vivo experiments are needed to determine the precise role of receptor exocytosis to the onset and maintenance of neurogenic inﬂammation. Therefore, regulation of TRPV1 surface density in nociceptor peripheral terminals could be an important mechanism for both the development and preservation of inﬂammatory hyperalgesia.

CONCLUDING REMARKS Regulated translocation of ion channels to the plasma membrane is a widespread strategy aimed at the spatiotemporal control of channel activity. Whether this sort of mechanism underlies the activation of TRP channels is still an open question that requires further documentation. However, it is becoming clear that agonist-induced exocytosis of integral membrane proteins is important to ﬁnely tune the cellular

328

TRP Ion Channel Function in Sensory Transduction

FIGURE 23.3 (Color ﬁgure follows p. 234.) TRPV1 is translocated to the cell surface by regulated exocytosis. (A) TRPV1 colocalizes in neuronal vesicles with the v-SNARE protein VAMP2, as evidenced from the immunocytochemistry.39 (B) BoNTA abrogates the PKCinduced translocation of TRPV1 to the plasma membrane, as concluded from biotinylation of surface-expressed receptors.39,40 TPA denotes 12-O-tetradecanoylphorbol-13-acetate, a potent agonist of PKC.

response to environmental signals. The number of channels and receptors that experience regulated exocytosis in vivo may increase in the near future, as this novel mechanism becomes general for regulating ion channel activity.

ACKNOWLEDGMENTS We thank all members of our group and colleagues of collaboration groups for their fundamental contributions to the results herein presented. We are indebted to ﬁnancial support from the MEC, GVA, Fundació La Caixa, and Fundación Ramón Areces.

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REFERENCES 1. Inoue, A. and Okabe, S. (2003) The dynamic organization of postsynaptic proteins: translocating molecules regulate synaptic function. Curr. Opin. Neurobiol. 13, 332–340. 2. Kavalali, E.T. (2002) SNARE interactions in membrane trafﬁcking: a perspective from mammalian central synapses. BioEssays 24, 926–936. 3. Buckley, K.M. et al. (2000) Regulation of neuronal function by protein trafﬁcking: a role for the endosomal pathway. J. Physiol. 525.1, 11–19. 4. Royle, S.J and Murrel-Lagnado, R.D. (2002) Constitutive cycling: a general mechanism to regulate cell surface proteins. BioEssays 25, 39–46. 5. Jahn, R., Lang, T., and Sudhof, T.C. (2003) Membrane fusion. Cell 112, 519– 533. 6. Watson, R.T., Kanzaki, M., and Pessin, J.E. (2004) Regulated membrane trafﬁcking of the insulin responsive glucose transporter 4 in adipocytes. Endocrine Rev. 25, 177–204. 7. Valenti, G. et al. (2005) Aquaporin 2 trafﬁcking. Endocrinology. Epub ahead of print. PMID: 16150901. 8. Peters, K.W. et al. (2001) Role of SNARE proteins in CFTR and ENaC trafﬁcking. Pﬂügers Arch. 443, S65–S69. 9. Barry, M.F. and Ziff, E.B. (2002) Receptor trafﬁcking and the plasticity of excitatory synapses. Curr. Opin. Neurobiol. 12, 279–286. 10. Montell, C. (2004) Exciting trips for TRPs. Nat. Cell. Biol. 6, 690–692. 11. Clapham, D.E. (2003) TRP channels as cellular sensors. Nature 426, 517–524. 12. Montell, C., Birnbaumer, L., and Flockerzi, V. (2002) The TRP channels, a remarkably functional family. Cell 108, 595–598. 13. Cosens, D.J. and Manning, A. (1969) Abnormal retinogram from a Drosophila mutant. Nature 224, 285–287. 14. Niemeyer, B.A. et al. (1996) The Drosophila light-activated conductance is composed of the two channels TRP and TRPL. Cell 85, 651–659. 15. Moran, M.M., Xu, H., and Clapham, D.E. (2004) TRP ion channels in the nervous system. Curr. Opin. Neurobiol. 14, 362–369. 16. Garcia-Sanz, N. et al. (2004) Identiﬁcation of a tetramerization domain in the Cterminus of the vanilloid receptor. J. Neurosci. 24, 5307–5314. 17. Phillips, A.M., Bull, A., and Kelly, L.E. (1992) Identiﬁcation of a Drosophila gene encoding a calmodulin-binding protein with homology to the trp phototransduction gene. Neuron 8, 631–642. 18. Chevesich, J., Kreuz, A.J., and Montell, C. (1997) Requirement for the PDZ domain protein, INAD, for localization of the TRP store-operated channel to a signaling complex. Neuron 18, 95–105. 19. Parekh, A.B. and Putney, J.W., Jr. (2005) Store-operated calcium channels. Physiol. Rev. 85, 757–810. 20. Hoth, M. and Penner, R. (1993) Calcium release–activated calcium current in rat mast cells. J. Physiol. 465, 359–386. 21. Venkatachalam, K. et al. (2002) The cellular and molecular basis of store-operated calcium entry. Nat. Cell Biol. 4, E263–E272. 22. Liu, X. et al. (2000). Trp1, a candidate protein for the store-operated Ca2+ inﬂux mechanism in salivary gland cells. J. Biol. Chem. 275, 3403–3411. 23. Yue, L. et al. (2001) CaT1 manifests the pore properties of the calcium release–activated calcium channel. Nature 410, 705–709.

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24. Schindl, R. et al. (2002) Store depletion–activated CaT1 currents in rat basophilic leukaemia mast cells are inhibited by 2-amino ethoxydiphenyl borate. Evidence for a regulatory component that controls activation of both CaT1 and CRAC (Ca2+ release–activated Ca2+ current) channels. J. Biol. Chem. 277, 26950–26958. 25. Voets, T. et al. (2001) CaT1 and the calcium release–activated calcium channel manifest distinct pore properties. J. Biol. Chem. 276, 477767–47770. 26. Kahr, H. et al. (2004) CaT1 knock-down fail to affect CRAC channels in mucosaltype mast cells. J. Physiol. 557, 121–132. 27. Yao, Y et al. (1999) Activation of store-operated Ca2+ currents in Xenopus oocytes requires SNAP-25 but not a diffusible messenger. Cell 98, 475–485. 28. Alderton, J.M. et al. (2000) Evidence for a vesicle-mediated maintenance of storeoperated calcium channels in a human embryonic kidney cell line. Cell Calcium 28, 161–169. 29. Scott, C.C. et al. (2003) Activation of store-operated calcium channels. Assessment of the role of SNARE-mediated vesicular transport. J. Biol. Chem. 278, 30534–30539. 30. Bahner, M et al. (2002) Light-regulated subcellular translocation of Drosophila TRPL channels induces long-term adaptation and modiﬁes the light-induced current. Neuron 34, 83–93. 31. Xu, Z.Z. and Sternberg, P.W. (2003) A C. elegans sperm TRP protein required for sperm–egg interactions during fertilization. Cell 114, 285–297. 32. Singh, B.B. et al. (2004) VAMP2-dependent exocytosis regulates plasma membrane insertion of TRPC3 channels and contributes to agonist-stimulated Ca2+ inﬂux. Molecular Cell 15, 635–646. 33. Cayouette, S. et al. (2004) Exocytotic insertion of TRPC6 channel into the plasma membrane upon Gq protein-coupled receptor activation. J. Biol. Chem. 279, 7241–7246. 34. Bezzerides, V. et al. (2004) Rapid vesicular translocation and insertion of TRP channels. Nat. Cell. Biol. 6, 709–720. 35. Greka, A. et al. (2003) TRPC5 is a regulator of hippocampal neurite length and growth cone morphology. Nat. Neurosci. 6, 837–845. 36. Kanzaki, M. et al. (1999) Translocation of a calcium-permeable cation channel induced by insulin-like growth factor-1. Nat. Cell. Biol. 1, 165–170. 37. Boels, K. et al. (2001) The neuropeptide head activator induces activation and translocation of the growth-factor-regulated Ca2+-permeable channel GRC. J. Cell Sci. 114, 3599–3606. 38. van de Graaf, S.F.J. et al. (2003) Functional expression of the epithelial Ca2+ channels (TRPV5 and TRPV6) requires association of the S100A10-annexin 2 complex. EMBO J. 22, 1478–1487. 39. Morenilla-Palao, C. et al. (2004) Regulated exocytosis contributes to protein kinase C potentiation of vanilloid receptor activity. J. Biol. Chem. 279, 25665–25679. 40. Van Buren, J.J. et al. (2005) Sensitization and translocation of TRPV1 by insulin and IGF-1. Mol. Pain. 1, 1–11. 41. Cui, M., Khanijou, S., and Aoki, K.R. (2004) Subcutaneous administration of botulinum toxin A reduces formalin-induced pain. Pain 107, 125–133.

24

Protein–Protein Interactions in TRPC Channel Complexes Petra Eder Karl Franzens University of Graz

Rainer Schindl University of Linz

Christoph Romanin University of Linz

Klaus Groschner Karl Franzens University of Graz

CONTENTS Abstract ..................................................................................................................332 Introduction............................................................................................................332 TRPC Proteins as Versatile Components of Native Ca2+ Signaling Complexes ........................................................332 Subunit Assembly and Anchoring of TRPC Channels within Multimolecular Signalplexes......................................................................333 Interactions Involved in Oligomerization and Assembly of Pore Complexes ....................................................................................333 Interactions Involved in Anchoring TRPC Channels within Multimolecular Signalplexes..........................................................335 Cellular Trafﬁcking of TRPC Channel Complexes ..............................................338 TRPC Channel Gating...........................................................................................339 Interactions Potentially Involved in Activation/Deactivation ...................340 Protein-Protein Interactions Potentially Involved in Inactivation ............................................................................................341 TRPC Signaling Partnerships ................................................................................342 Concluding Remarks and Perspectives .................................................................342

331

332

TRP Ion Channel Function in Sensory Transduction

Appendix: Abbreviations .......................................................................................343 References..............................................................................................................343

ABSTRACT Screening for homologues of the Drosophila trp (transient receptor potential) gene product has uncovered a large family of membrane proteins of which the closest relatives to the Drosophila protein have been assigned to the canonical or classical subfamily (TRPC). The prominent physiological function of these proteins, as delineated from heterologous expression and knockdown experiments in native cells, appears to be sensing phospholipase C (PLC)–derived stimuli and conversion of this input into cellular Ca2+ signals. Another common feature of TRPC proteins is the ability to form cation-conducting pore structures. Thus, the role of TRPCs in PLCdependent Ca2+ signaling has been attributed to the formation of regulated Ca2+ entry channels. Despite the existence of this unifying principle of TRPC signal transduction, an unforeseen complexity of the roles of TRPC proteins within the cellular Ca2+ signaling network emerged. The contribution of TRPC proteins to cellular Ca2+ homeostasis may involve formation of a wide variety of different Ca2+ entry pathways that contain distinct TRPC homo-or heteromultimeric pore structures, along with a multitude of regulatory proteins and scaffolds. Here, we summarize current knowledge on protein–protein interactions that are of potential signiﬁcance for the formation and function of native TRPC channel complexes and highlight recent concepts regarding the role of these interactions for cellular control of plasma membrane cation conductances and cellular Ca2+ signaling.

INTRODUCTION TRPC PROTEINS AS VERSATILE COMPONENTS OF NATIVE CA2+ SIGNALING COMPLEXES The contribution of TRPC proteins to PLC-dependent Ca2+ entry has been demonstrated in a variety of cell lines and native cells by antisense, siRNA, and dominant negative knockdown experiments.27,30,40,41,65–67 Three lines of evidence suggest that most native TRPC Ca2+ entry channels are heteromeric, multisubunit complexes embedded into large signalplexes composed of scaffolds, regulatory proteins, and speciﬁc signaling partners, which determine the Ca2+ entry pathway’s biophysical, pharmacological, and regulatory properties. First, antisense and siRNA as well as dominant negative knockdown strategies provided strong evidence for contribution of multiple TRPC proteins in the permeation pathway of endogenous phospholipase– regulated cation channels.30,65–67 Second, overexpression of a particular TRPC species into a different background of regulatory proteins or potential heteromerization partners was found to promote or generate distinctly different Ca2+ entry phenomena,28,50,58 and the reconstitution of native cation conductances by overexpression of a single TRPC species in standard expression systems was barely successful. Third, a multitude of protein interaction partners with potential impact on TRPC channel localization and gating has been identiﬁed. Consequently, the concept of

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333

TRPC proteins as multifunctional components of Ca2+ signaling complexes has emerged, and analysis of TRPC protein–protein interactions has been recognized as a key step to understanding the physiological role of these proteins. A number of different functional aspects may be considered for protein–protein interactions within ion channel signalplexes. Accordingly, we aim to provide an overview on current knowledge regarding TRPC protein–protein interactions in view of their potential signiﬁcance for (1) assembly, subunit composition of the pore complex, and its anchoring within larger, multimolecular signalplexes; (2) cellular trafﬁcking of channel complexes; (3) channel activation and inactivation and terms of gating processes; and (4) functional interaction with other signaling components. Inevitable overlaps between these functional aspects of protein–protein interactions will be highlighted and discussed.

SUBUNIT ASSEMBLY AND ANCHORING OF TRPC CHANNELS WITHIN MULTIMOLECULAR SIGNALPLEXES The Drosophila TRP protein has been recognized as a multifunctional component of the signalplex that enables transduction of light into a photoreceptor Ca2+ signal (for review, see reference 37). Two types of protein–protein interactions may be considered for the formation of functional TRP signalplexes (i.e., interactions that stabilize the pore-forming TRP oligomer and interactions that anchor this pore structure to scaffolds and regulatory components).

INTERACTIONS INVOLVED COMPLEXES

IN

OLIGOMERIZATION

AND

ASSEMBLY

OF

PORE

TRP proteins contain six transmembrane domains, a pore-loop motif between the ﬁfth and sixth domain, and a cytosolic N- and C-terminus (Figure 24.1). Functional TRP channels consist of a tetrameric structure, and heterologous expression results predominantly in homomultimeric assemblies. Nonetheless, heteromeric TRP channels have also been identiﬁed for the TRPC, TRPV, TRPM, and TRPP subfamilies, as well as for Drosophila TRPs. Heteromerization of mammalian TRPC channels has been clearly demonstrated among closely related members of subfamilies such as TRPC1/4/5 and TRPC3/6/7.18,50 Furthermore, TRPC1 seems to bear a speciﬁc potential for heteromerization or facilitation of heteromerization among distantly related TRPCs, as functional as well as physical interaction between TRPC1 and TRPC3 has been observed28,30 whereby the interaction domain was detected in the N-termini of the two channel proteins, and evidence for the existence of TRPC1/3/551 and TRPC1/3/773 heteromers has been provided. Moreover, the only reported example of an interaction between subunits of distinct subfamilies is that of TRPC1 and TRPP2, which involves the C-termini of these proteins. So far, the functional properties of these complexes and their physiological signiﬁcance have not been delineated.55 Heteromerization increases the variability of TRP channel architecture and function, resulting in TRPC1/5, TRPC1/4, or TRPC1/3/5 heteromers in unique permeation

334

TRP Ion Channel Function in Sensory Transduction

FIGURE 24.1 Structural features of TRPC proteins involved in protein–protein interactions. All TRPC family members, including Drosophila TRP, consist of a cytosolic N-terminus with a varying number of ankyrin repeats (Ank), a coiled-coil domain (CC), a transmembrane part composed of six transmembrane-spanning domains (dark boxes), and a cytosolic C-terminus encompassing the TRP box (EWKFAR), a proline-rich sequence (PP), the CaM- and IP3Rbinding site (CIRB), and a coiled coil domain (CC). A second CaM-binding site (CCB) has been located in the C-termini of TRPC 1, 2, 4, and 5. All domain structures refer to human sequences, except for TRPC2 (mouse) and Drosophila TRP.

properties. Such heteromultimers are likely expressed in native tissues as demonstrated for rat brains.50, 51 Interestingly, the cytosolic N-terminal strand has been suggested as an important interaction domain for various TRP channels like Drosophila TRPs,69,70 TRPC1,12,30 TRPC3,15 TRPM1,71 and TRPV6.9,13,21 Conserved potential interaction domains within the N-terminal strand are the ankyrin-like repeats and the coiled-coil domain. For TRPC1 and Drosophila TRP, the coiled-coil region was found essential for

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335

channel assembly and function,12,69 whereas studies with TRPV proteins highlighted the ankyrin-like repeats for TRPV5 and TRPV6 assembly.9,13 Moreover, TRPC3 mutants lacking the ankyrin-like repeat domain were found to lack channel function.62 Nonetheless the requirement of a stable, ion-selective pore suggests additional interaction motifs within the transmembrane domain. It may turn out that distinct assembly domains within one TRP protein may form either homo- or heteromeric assemblies.

INTERACTIONS INVOLVED IN ANCHORING TRPC CHANNELS MULTIMOLECULAR SIGNALPLEXES

WITHIN

TRPC pore complexes are linked to other signaling molecules via classical adapter proteins, resulting in assembly and stabilization of multimolecular signaling complexes.10,19,36,43,53,56 Besides their function as backbone scaffolds, which enable signaling molecules to cooperate in terms of an efﬁcient signal transduction unit, these adaptor proteins are likely to determine functions of individual proteins of the complex by generating a speciﬁc microenvironment or by direct regulatory effects. Thus, proteins that are considered in this section as essential scaffolds in TRPC complexes will be discussed below in view of their potential impact on functions other than mere complex assembly. For the Drosophila TRP channel, the multivalent PDZ scaffold protein INAD (inactivation no after potential D; Table 24.1) serves as the backbone structure that assembles a protein complex including the cation channel and a number of other signaling proteins such as the PLC (norpA) involved in channel activation and a regulatory PKC.19,26 These signaling components appear to be constitutively attached to the PDZ scaffold. Among the ﬁve PDZ domains in INAD, PDZ3 binds to the C terminus of TRPC channel protein,26 and this interaction is considered essential for retention of the TRPC channels in the rhabdomers of the photoreceptors.25,26 The TRPC/INAD interaction represents an example for mutual anchoring, as disruption of this interaction leads to mislocalization of INAD and the entire core complex.26 Hence, the Drosophila TRP protein is most likely a multifunctional player and has to be considered by itself as a scaffold of the signalplex. For mammalian TRPC4 and TRPC5 proteins, members of another family of PDZ domain scaffolds, NHERFs (sodium-hydrogen exchanger regulatory factors), appear to organize TRPC channel complexes.36,53 NHERF proteins are multifunctional scaffolds that contain two tandem PDZ protein interaction domains along with an ERM (ezrin, radixin, moesin, merlin) domain.60 NHERF1, the ezrin-binding phosphoprotein-50 (EBP50), has been identiﬁed as an adaptor that tethers TRPC4 channels to F-actin, resulting in a promotion of channel density in or close to the plasma membrane.36 The interaction between EBP50 and TRPC4/5 involves a domain in the cytoplasmic C-terminus of these TRPC channel proteins, which contains a “VTTRL” motif (Figure 24.1).36,38 NHERF proteins were recently found to associate with G-protein-coupled receptors, heterotrimeric G proteins, and receptor tyrosine kinases as recently reviewed in reference 63, as well as phospholipase Cβ.53 EBP50 is likely to organize multimolecular TRPC signaling units in mammalian cells. Besides NHERF1 (EBP50), which may play a role for the TRPC4/5

CaBP1 (22) CaM (76) Caveolin-1 (8) NHERF (36,38) Homer (72) Immunophilins (46) INAD (26) IP3R (52) Junctate (48,54) NCX (42) PLCy (57) RhoA (35) Stathmin (14) VAMP2 (45) ZO-1 (47)

Proposed Binding Domain

TRPC5 C-terminus TRPC1–7 CIRB; CCB (TRPC1,2,4,5; Drosophila TRP) TRPC1,3 TRPC1: aa271–349 TRPC4 +ß; TRPC5 C-terminus TRPC1,2,5 TRPC1: proline-rich domain+ N-terminus FKBP12: Drosophila TRP, TRPC3,6; FKBP52: TRPC1,4,5 ﬁrst dipeptide of proline-rich domain Drosophila TRP C-terminus TRPC1–7; Drosophila TRP CIRB TRPC2,3,5 TRPC2: C-terminus TRPC3 C-terminus TRPC3 TRPC3: aa 40–48 TRPC1 not determined TRPC5 TRPC5: coiled-coil domain in N-terminus TRPC3 TRPC3:aa123–221 TRPC4 C-terminus

Proposed TRPC Binding Partners

TABLE 24.1 Binding Partners of TRPC Channels and Their Effects on Channel Localization and Function

inactivation inactivation of TRPC trafﬁcking and anchoring anchoring activation/inactivation not determined anchoring conformational coupling and activation activation increased NCX mediated Ca2+ entry targeting, anchoring trafﬁcking trafﬁcking and targeting trafﬁcking targeting

Functional consequence

336 TRP Ion Channel Function in Sensory Transduction

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channel complex organization in neuronal tissues, NHERF2 has recently been suggested as a scaffold of TRPC4 channels in the endothelium.25 Similarly, another PDZ-scaffolding protein, zonula occludens 1 (ZO-1), has been reported to interact with TRPC4 in astrocytes.47 TRPC-mediated signal transduction in mammalian cells has been suggested to take place in complex assemblies that are localized in membrane subdomains.1,2 Like many other signaling molecules, TRPC proteins appear to be sequestered and segregated from other membrane constituents by accumulation in membrane domains characterized by a speciﬁc lipid composition and physical properties termed lipid rafts.2,16 Thus the molecular organization of TRPC signaling complexes in mammalian cells may involve interplay between lipid- and protein-mediated assembly of multimolecular complexes. TRPCs, speciﬁcally TRPC1 and TRPC3, have been identiﬁed as components of signalplexes that reside in a speciﬁc type of lipid microdomain termed caveolae.2,31,32 Organization of signaling assemblies in caveolae involves caveolins, a family of raft-stabilizing scaffold proteins, which serve as adaptor and regulator proteins and as an interface between membrane proteins and lipids.11 The prototypical family member, caveolin-1, is a palmitoylated protein with a unique hairpin structure ﬂanked by two membrane attachment domains, one of which includes a scaffold domain (CSD; aa 92–101) mediating protein–protein interactions. TRPC proteins contain a putative caveolin binding domain in the cytoplasmic N-termini, which has clearly been localized for TRPC1 (aa 271–349; Figure 24.1, Table 24.1)8 and which is expected to associate with the CSD, thereby tethering TRPC channels to caveolin-1 and caveolar signalplexes. Thus, at least some TRPC channel complexes appear to reside within a speciﬁc membrane lipid environment, which may be essential for proper organization of the signaling complexes or cellular regulation of channel activity. Speciﬁcally, TRPC1-containing channel complexes that are controlled by the ﬁlling state of intracellular Ca2+ stores (storeoperated Ca2+ entry channels, SOCs) have been suggested as components of caveolar signalplexes.32 Caveolin-1 is not only a scaffold protein that stabilizes lipid microdomains and anchors a variety of signaling molecules to these domains, but it also serves as an inhibitory regulator that sequesters signaling molecules within the plasma membrane in an inactive, resting state.11,64 Moreover, caveolins are important players in vesicular transport processes, and the role of caveolin–TRPC interactions in trafﬁcking and regulation of TRPC channels is incompletely understood. It remains to be clariﬁed if the caveolin–TRPC association is simply stabilizing channel complexes of distinct regulatory properties or if caveolin–TRPC complexes correspond to a speciﬁc functional state of TRPC channels (resting, activated, inactivated). As vesicular trafﬁcking of mammalian TRPC channel complexes has been suggested as a key mechanism of TRPC conductance activation and inactivation via rapid insertion and retrieval of channel complexes, a role of caveolin in this mechanism may be considered. In general, combined protein–protein and protein–lipid interactions appear as a key principle for assembly of functional TRPC complexes in the plasma membrane. Besides association with the lipid-binding scaffold caveolin, TRPC3 was found to associate with PLCγ1 to form a unique, composite PH domain, which binds PIP2 as well as sphingosine-1-phosphate.57 The lipid-binding properties of this bimolecular

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domain, which is formed by association of two incomplete lipid-binding domains residing in the very N-terminus of TRPC3 (Figure 24.1) and in PH-c of PLCγ1, were found essential for plasma membrane targeting and function of TRPC3 channel complexes.57 Again, it remains to be clariﬁed if the impact of this complex interaction on channel function is based merely on the stabilization of a plasma membrane– associated TRPC channel complex or if an additional regulatory principle is enabled by the interaction.

CELLULAR TRAFFICKING OF TRPC CHANNEL COMPLEXES Analysis of the functional role of the TRP–INAD interactions revealed that the TRP–INAD association is crucial for anchoring and retention of signalplexes in the rhabdomers but not for targeting of the TRP protein itself.37,56 Interestingly, mutational impairment of the direct TRP–INAD interaction barely affects the light response of the photoreceptors,26 while a profound alteration in regulatory channel properties (i.e., a gain in the ability to recognize Ca2+ store depletion as an activating input stimulus) was detected as a consequence of TRPC–INAD uncoupling.17 Drosophila TRP channels appear to be correctly targeted even without direct attachment to the backbone scaffold INAD and do not display marked dynamic trafﬁcking during light stimulation. By contrast, TRPL—a potential component of the TRP transducisome—displays profound, light-induced translocation that is considered a key event in adaptation of the photoreceptors.3 The molecular mechanism of this dynamic trafﬁcking of TRPL is still elusive. Nonetheless, the presence of TRP regardless of its ion channel function is essential for light-dependent shuttling of TRPL, indicating a nonchannel, structural role of TRP and a possible plasticity of the subunit stoichiometry involving insertion and retrieval of a channel subunit. Similar dynamic trafﬁcking has recently been reported for mammalian TRPC proteins. There is a fairly general agreement to consider mammalian TRPC proteins as the basis of receptor-PLC-regulated cation conductances including also some speciﬁc store-operated Ca2+ conductances. Cellular regulation of such TRPC conductances is still enigmatic. Rapid insertion and retrieval of channel complexes in the plasma membrane has recently emerged as a potential process involved in regulating TRPC conductances. A recent study on the cellular trafﬁcking of TRPC5 channels using evanescent ﬁeld microscopy revealed rapid shuttling of TRPC channels between a speciﬁc, closely membrane–associated vesicle pool and the plasma membrane.5 Although these results demonstrate efﬁcient and rapid transport of channels toward the plasma membrane in response to stimuli that activate the TRPC conductance, it remains elusive whether rapid vesicular insertion of channels without additional gating processes is sufﬁcient to explain conductance activation. Nonetheless, vesicular trafﬁcking represents a potentially important mechanism that determines the availability of functional TRPC complexes in the membrane. Several protein–protein interactions appear important for such translocation and insertion-retrieval processes. TRPC5 was found to associate with neuron-speciﬁc members of the stathmin family of phosphoproteins.14 Stathmins are important

Protein–Protein Interactions in TRPC Channel Complexes

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determinants of microtubule organization and are highly expressed during neuronal development.49 Association of TRPC5 with stathmin 2 involves the classical α-helical stathmin domain in the protein’s C-terminus and a coiled-coil domain in the N-terminus of TRPC5 (Figure 24.1).14 This interaction loads neuronal TRPC5 to transport vesicles that are targeted to growth cones. Importantly, correct targeting of stathmin complexes to vesicles requires a speciﬁc protein–lipid interaction, as palmitoylation of stathmin is essential for its accumulation in vesicles.33 The stathmin-associated, neuronal TRPC5 channel may reside in a complex, containing synaptic vesicle proteins such as VAMP2 and synaptotagmin, as indicated by colocalization of these proteins in developing hippocampal neurons.14 For TRPC3, association with SNARE proteins has been demonstrated in both the HEK293 expression system as well as in native tissues.45 Similar to TRPC5, TRPC3 channels have been shown to reside in a population of mobile, sub-plasmalemmal vesicles, from which the protein is delivered toward the plasma membrane during stimuli that activated cellular TRPC3 conductances. The cytoplasmic N-terminus of TRPC3 was found to associate with VAMP2 and αSNAP, and an interaction domain responsible for VAMP2 binding has been identiﬁed (aa 123–221 in TRPC3; Figure 24.1).45 This putative VAMP2 interaction motif is present also in the closest relative of TRPC3, TRPC6. A variety of other SNARE proteins such as syntaxin, SNAP23, and NSF were found to colocalize with TRPC3 channels in the HEK293 expression system, and endogenous neuronal TRPC3 was co-immunoprecipitated with both VAMP2 and syntaxin 3, substantiating the existence of native TRPC3–SNARE complexes. Thus, rapid vesicular trafﬁcking of TRPC channels involves association of the pore complex with an array of vesicle-targeted proteins. Dynamic vesicular transport and targeting of TRPC channels are likely to involve complex protein–lipid interactions as indicted by the essential role of lipid anchoring of complex components such as stathmin14 and by the recognized key role of lipid-, speciﬁcally PIP2, metabolism for rapid vesicular trafﬁcking.5 Protein–protein interactions that are primarily considered to promote or enable targeting and stabilization of signal complexes in membrane domains, such as NHERFs and caveolins, in dynamic, regulated vesicular transport appear likely but need to be delineated in further studies. Similarly, an overlap may exist between regulatory protein–protein interactions that are of potential importance for gating processes in TRPC channels and dynamic trafﬁcking of channel complexes. Moreover, targeting mechanisms may involve essential, secondary proteins, such as RhoA, a monomeric GTP-binding protein, which has been shown to associate with IP3R and TRPC1 upon stimulation with thrombin. Membrane insertion of this complex triggers Ca2+ entry through TRPC1 after store depletion, which may be involved in endothelial permeability.35

TRPC CHANNEL GATING Besides insertion and retrieval of constitutively conducting TRPC pore complexes as a principle of conductance regulation, classical gating of native TRPC channel complexes by either second messenger or interaction with regulatory proteins is

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considered as well. Such gating processes may be the basis of activation/deactivation as well as of inactivation of TRPC-related membrane conductances.

INTERACTIONS POTENTIALLY INVOLVED

IN

ACTIVATION/DEACTIVATION

TRPC channel complexes that are controlled by the ﬁlling state of intracellular Ca2+ storage compartments have been postulated to communicate with these storage compartments via protein–protein interactions involving complex partners that are part of intracellular membrane structures.6,54,75 This gating principle has been termed as a conformational coupling model4, 20 for store-operated activation and deactivation of TRPC-related Ca2+ entry channels. A TRPC species that has repeatedly been implicated in the formation of native SOCs is TRPC3. One candidate mediating communication of signals from the ER to the plasma membrane TRPC channel is the IP3R, which has been proposed to play a central role in SOC activation. The model of a tight functional link between SOCs and the IP3R is based on studies demonstrating an inhibitory effect of the IP3R antagonist 2-APB (2-aminoethyl diphenylborate) on PLC-mediated TRPC activation34 and the ability of activated IP3R to interact and regulate TRPC3.24 The TRPC3 site for association with the IP3R, which by itself contains two TRPC-interacting sequences, has been located between aa 742–795.6 IP3Rs were also found to interact with TRPC1 and TRPC6,6,32 substantiating the potential signiﬁcance of IP3R–TRP interactions as a gating mechanism for TRP channels. The regulatory TRPC–IP3R interaction is governed by Ca2+ calmodulin (CaM), which has been shown to compete with the IP3R binding site of any TRPC.52,75 Thus, this speciﬁc binding site has been termed a CaM- and IP3R-binding site (CIRB) (Figure 24.1) and is highly conserved among TRPC channels, including also the three Drosophila proteins.76 In terms of a conformational coupling model, the activated IP3R is supposed to displace CaM from CIRB (aa 761–795), representing a crucial step in activating TRPC3 channels. Alternatively, the CIRB region has been recognized as a structure essential for correct channel targeting as deletions within this region impair TRPC3 function at least in part by preventing its targeting to the plasma membrane,62 and the CIRB domain was suggested to govern membrane localization and targeting independent of an interaction with IP3R or CaM.62 Information transfer between intracellular membranes and plasma membrane TRPCs in terms of conformational coupling may in addition to IP3Rs involve regulatory proteins of the ER. Homer proteins and junctate are candidates regulating the interaction between TRPC and the IP3R. Encoded by three gene families (Homer 1–3), Homer proteins comprise an EVH1 domain (enabled/vasodilator-stimulated phosphoprotein homology 1), which binds proline-rich sequences and a coiled-coil domain, which is necessary for self-association.68 Upon interaction with prolinerich motifs, Homer crosslinks proteins into a macromolecular complex, as shown for metabotropic glutamate receptors7 and a series of other proteins. Because TRPC and IP3R exhibit proline-rich sequences, a possible link between these proteins and Homer has been considered. Indeed, Homer interacts with the N-terminus of IP3R and the proline-rich sequence in the C-terminus, as well as the N-terminus of TRPC1 under resting conditions (Figure 24.1).72 Following receptor stimulation or store depletion, the complex disassembles with an increase of TRPC1 activity. The

Protein–Protein Interactions in TRPC Channel Complexes

341

TRPC1–Homer–IP3R disassembly is regulated by Homer 1a, which lacks the coiledcoil domain and dissociates Homer complexes upon agonist stimulation. In case of the TRPC1–Homer–IP3R complex, disruption by Homer 1a renders TRPC1 in a spontaneously active state unresponsive to further stimulation. The physiological relevance of the regulative impact of Homer on the TRPC–IP3R complex has been demonstrated in human salivary gland cells where TRPC1 is part of the SOC. It is obvious to speculate that further signaling proteins intervene in this process. Immunophilins, for example, bind speciﬁcally and selectively to TRPC1, 3–7 (FKBP12, FKBP52) via the LP dipeptide located in the C-terminal proline-rich sequence (Figure 24.1)46 and could consequently interfere with TRPC1–Homer interaction. Junctate is another protein that has attracted attention as an important regulatory protein involved in IP3R-dependent Ca2+ inﬂux.54 Junctate is an integral protein of the endo/sarcoplasmic membrane consisting of a short N-terminal cytoplasmic domain interacting with the IP3R, followed by a hydrophobic transmembrane domain and a luminal C-terminal calcium-binding domain. Recently, members of the TRPC family (TRPC2, 3, and 5) have been identiﬁed to build up a supramolecular complex together with IP3R and junctate.48,54 A binding site for junctate has been proposed in the C-terminus of TRPC3 (Figure 24.1). The functional relevance of such complexes has been studied in heterologous expression systems and rodent sperm.48,54 In HEK293 cells, junctate induces an increase in the Ca2+ releasable from intracellular stores. This effect involves the interaction between IP3R and junctate and its capability to bind Ca2+. It has been hypothesized that IP3R association with junctate results in enhanced TRPC-mediated Ca2+ entry. Two principles are considered responsible for this promotion of TRPC activation: (1) receptor activation triggers junctate binding to both the IP3R and the TRPC channel, resulting in an increased Ca2+ entry; (2) Ca2+ entry is initiated by a reduction of Ca2+ at the luminal Ca2+binding domain of junctate.54 Interestingly, another ER protein, designated as STIM, contains a luminal Ca2+-binding domain and was able to sense the ﬁlling state of the ER.29 Reduced luminal Ca2+ binding in response to store depletion triggered STIM translocation sites near or in the plasma membrane,74 and the protein was demonstrated as an essential component of store-operated Ca2+ inﬂux mechanisms. Thus, it appears of particular interest to examine the role of STIM in TRPC-mediated, store-operated Ca2+ entry.

PROTEIN-PROTEIN INTERACTIONS POTENTIALLY INVOLVED

IN INACTIVATION

Inactivation of TRPC channels has been well studied and is mainly determined on coaction of Ca2+ and calmodulin (CaM). All TRPC members include in their C-termini a competitive, Ca2+-dependent site for CaM and IP3 receptor binding (CIRB). Whereas low Ca2+ concentrations favor IP3R binding, higher Ca2+ levels ranging from 10 nM to 290 nM for certain TRPCs promote CaM binding,52 which is considered to terminate activation via reassociation with IP3R. Indeed, electrophysiological measurements demonstrated that an N-terminal peptide of the IP3R directly activates single TRPC3 or TRPC4 channels, while displacement of this peptide by CaM inhibited the channel,23,75 suggesting these ion channels as candidates for the conformational coupling model for store-operated channels.

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TRP Ion Channel Function in Sensory Transduction

TRP channels include both N-terminal and C-terminal CaM-binding (CCB) sites.76 Evidence for functional signiﬁcance of the N-terminal CaM-binding domain is so far lacking. The CCB domain of TRPC1, located at the very end of its C-terminus and overlapping with the coiled-coil domain,59 mediates Ca2+/CaM-dependent inactivation (Figure 24.1).44 By contrast, the same domain has recently been shown to enhance activity of TRPC5.39 Inhibition of this ion channel has been observed, as the results of an interaction with the Ca2+-binding protein 1 (CaBP1).22 Unlike CaM, which is ubiquitously expressed, CaBP1 is speciﬁcally expressed in neurons and binds to the C-terminus of TRPC5 (Figure 24.1, Table 24.1), close to the ﬁrst CCB site competing with CaM,22 suggesting ﬁne tuning of channel activity by these Ca2+-binding proteins.

TRPC SIGNALING PARTNERSHIPS Protein–protein interactions are considered to determine TRPC channel membrane targeting and function as outlined above. Besides control of TRPC function as storeoperated or receptor-operated channels, speciﬁc protein–protein interactions may link TRPC channels to other Ca2+ transport systems, which function as signaling partners that generate distinct spatial and temporal Ca2+ signals. TRPC channels have been demonstrated to interact functionally with voltage-gated calcium channels41 or the sodium calcium exchanger (NCX). A functional link between NCX (CalX) and TRP channels has recently been found in Drosophila. Here it seems that an increased Ca2+ entry mediated by TRP is compensated by enhanced NCX activity extruding Ca2+, which is considered to prevent ischemic cell death.61 Colocalization and functional interaction between NCX and TRP has been detected in rhabdomers.37 NCX–TRPC complex formation has also been suggested from results obtained in heterologous expression studies with TRPC3.42 Na+ entry mediated by TRPC3 has been shown to enable local accumulation of Na+ sufﬁcient to drive Ca2+ entry via reverse mode NCX. This functional interaction is underlined by a tight physical interaction between TRPC3 and NCX1 (Table 24.1), suggesting a close spatial proximity between these ion transport systems.

CONCLUDING REMARKS AND PERSPECTIVES TRP proteins have emerged as multifunctional players in signalplexes essential for cellular Ca2+ signaling. The high versatility of TRP and speciﬁcally of TRPC proteins to serve as key components of a variety of Ca2+-permeable cation channels with distinct regulatory and biophysical properties as well as the striking ability of TRP channel complexes to integrate multiple input stimuli is apparently based on a high complexity of protein–protein interactions in TRP channel complexes. Such signaling complexes may be viewed as paradigm “channelosomes” that govern cell functions in a highly ﬂexible manner to allow tissue- and cell state– speciﬁc cellular responses. Further analysis of the protein–protein interactions in TRP channelosomes may uncover avenues toward using TRPC Ca2+ channels as therapeutic targets.

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APPENDIX: ABBREVIATIONS CaBP1 Ca2+ binding protein 1 CaM calmodulin CCB C-terminal CaM binding site CIRB CaM-and IP3R- binding site EBP50 ezrin-binding phosphoprotein-50 ER endoplasmic reticulum EVH1 enabled/vasodilator-stimulated phosphoprotein homology1 INAD inactivation no after potential D IP3 inositol trisphosphate IP3R IP3 receptor NHERF Na+/H+ exchange regulatory factor NSF N-ethylmaleimide sensitive factor PKC protein kinase C PLC phospholipase C PM plasma membrane ROC receptor-operated channel SNAP soluble N-ethylmaleimide-sensitve factor attachment protein SNARE soluble N-ethylmlaleimide-senitive factor attachment protein receptor SOC store-operated channel STIM stromal interaction molecule TRPC/V/M/PP channel canonical/vanilloid/melastatin/polycystin transient receptor potential channel VAMP vesicle-associated membrane protein

REFERENCES 1. Ambudkar IS. Cellular domains that contribute to Ca2+ entry events. Sci STKE 2004:pe32, 2004. 2. Ambudkar IS, Brazer SC, Liu X, Lockwich T, and Singh B. Plasma membrane localization of TRPC channels: role of caveolar lipid rafts. Novartis Found Symp 258:63–70; discussion 70–64, 98–102, 263–306, 2004. 3. Bahner M, Frechter S, Da Silva N, Minke B, Paulsen R, and Huber A. Light-regulated subcellular translocation of Drosophila TRPL channels induces long-term adaptation and modiﬁes the light-induced current. Neuron 34:83–93, 2002. 4. Berridge MJ. Capacitative calcium entry. Biochem J 312 (Pt. 1):1–11, 1995. 5. Bezzerides VJ, Ramsey IS, Kotecha S, Greka A, and Clapham DE. Rapid vesicular translocation and insertion of TRP channels. Nat Cell Biol 6:709–720, 2004. 6. Boulay G, Brown DM, Qin N, Jiang M, Dietrich A, Zhu MX, Chen Z, Birnbaumer M, Mikoshiba K, and Birnbaumer L. Modulation of Ca(2+) entry by polypeptides of the inositol 1,4,5-trisphosphate receptor (IP3R) that bind transient receptor potential (TRP): evidence for roles of TRP and IP3R in store depletion-activated Ca(2+) entry. Proc Natl Acad Sci USA 96:14955–14960, 1999. 7. Brakeman PR, Lanahan AA, O'Brien R, Roche K, Barnes CA, Huganir RL, and Worley PF. Homer: a protein that selectively binds metabotropic glutamate receptors. Nature 386:284–288, 1997. 8. Brazer SC, Singh BB, Liu X, Swaim W, and Ambudkar IS. Caveolin-1 contributes to assembly of store-operated Ca2+ inﬂux channels by regulating plasma membrane localization of TRPC1. J Biol Chem 278:27208–27215, 2003.

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9. Chang Q, Gyftogianni E, van de Graaf SF, Hoefs S, Weidema FA, Bindels RJ, and Hoenderop JG. Molecular determinants in TRPV5 channel assembly. J Biol Chem 279:54304–54311, 2004. 10. Chevesich J, Kreuz AJ, and Montell C. Requirement for the PDZ domain protein, INAD, for localization of the TRP store-operated channel to a signaling complex. Neuron 18:95–105, 1997. 11. Cohen AW, Hnasko R, Schubert W, and Lisanti MP. Role of caveolae and caveolins in health and disease. Physiol Rev 84:1341–1379, 2004. 12. Engelke M, Friedrich O, Budde P, Schafer C, Niemann U, Zitt C, Jungling E, Rocks O, Luckhoff A, and Frey J. Structural domains required for channel function of the mouse transient receptor potential protein homologue TRP1beta. FEBS Lett 523:193–199, 2002. 13. Erler I, Hirnet D, Wissenbach U, Flockerzi V, and Niemeyer BA. Ca2+-selective transient receptor potential V channel architecture and function require a speciﬁc ankyrin repeat. J Biol Chem 279:34456–34463, 2004. 14. Greka A, Navarro B, Oancea E, Duggan A, and Clapham DE. TRPC5 is a regulator of hippocampal neurite length and growth cone morphology. Nat Neurosci 6:837–845, 2003. 15. Groschner K, Hingel S, Lintschinger B, Balzer M, Romanin C, Zhu X, and Schreibmayer W. Trp proteins form store-operated cation channels in human vascular endothelial cells. FEBS Lett 437:101–106, 1998. 16. Harder T. Formation of functional cell membrane domains: the interplay of lipid- and protein-mediated interactions. Philos Trans R Soc Lond B Biol Sci 358:863–868, 2003. 17. Harteneck C, Kuchta SN, Huber A, Paulsen R, and Schultz G. The PDZ scaffold protein INAD abolishes apparent store-dependent regulation of the light-activated cation channel TRP. Faseb J 16:1668–1670, 2002. 18. Hofmann T, Schaefer M, Schultz G, and Gudermann T. Subunit composition of mammalian transient receptor potential channels in living cells. Proc Natl Acad Sci USA 99:7461–7466, 2002. 19. Huber A, Sander P, Gobert A, Bahner M, Hermann R, and Paulsen R. The transient receptor potential protein (Trp), a putative store-operated Ca2+ channel essential for phosphoinositide-mediated photoreception, forms a signaling complex with NorpA, InaC, and InaD. Embo J 15:7036–7045, 1996. 20. Irvine RF. 'Quantal' Ca2+ release and the control of Ca2+ entry by inositol phosphates— a possible mechanism. FEBS Lett 263:5–9, 1990. 21. Kahr H, Schindl R, Fritsch R, Heinze B, Hofbauer M, Hack ME, Mortelmaier MA, Groschner K, Peng JB, Takanaga H, Hediger MA, and Romanin C. CaT1 knockdown strategies fail to affect CRAC channels in mucosal-type mast cells. J Physiol 557:121–132, 2004. 22. Kinoshita-Kawada M, Tang J, Xiao R, Kaneko S, Foskett JK, and Zhu MX. Inhibition of TRPC5 channels by Ca(2+)-binding protein 1 in Xenopus oocytes. Pﬂügers Arch 450:345–354, 2005. 23. Kiselyov K, Mignery GA, Zhu MX, and Muallem S. The N-terminal domain of the IP3 receptor gates store-operated hTrp3 channels. Mol Cell 4:423–429, 1999. 24. Kiselyov K, Xu X, Mozhayeva G, Kuo T, Pessah I, Mignery G, Zhu X, Birnbaumer L, and Muallem S. Functional interaction between InsP3 receptors and store-operated Htrp3 channels. Nature 396:478–482, 1998 25. Lee-Kwon W, Wade JB, Zhang Z, Pallone TL, and Weinman EJ. Expression of TRPC4 channel protein that interacts with NHERF-2 in rat descending vasa recta. Am J Physiol Cell Physiol 288:C942–949, 2005.

Protein–Protein Interactions in TRPC Channel Complexes

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26. Li HS and Montell C. TRP and the PDZ protein, INAD, form the core complex required for retention of the signalplex in Drosophila photoreceptor cells. J Cell Biol 150:1411–1422, 2000. 27. Li Y, Jia YC, Cui K, Li N, Zheng ZY, Wang YZ, and Yuan XB. Essential role of TRPC channels in the guidance of nerve growth cones by brain-derived neurotrophic factor. Nature 434:894–898, 2005. 28. Lintschinger B, Balzer-Geldsetzer M, Baskaran T, Graier WF, Romanin C, Zhu MX, and Groschner K. Coassembly of Trp1 and Trp3 proteins generates diacylglyceroland Ca2+-sensitive cation channels. J Biol Chem 275:27799–27805, 2000. 29. Liou J, Kim ML, Heo WD, Jones JT, Myers JW, Ferrell JE, Jr., and Meyer T. STIM is a Ca2+ sensor essential for Ca2+-store-depletion-triggered Ca2+ inﬂux. Curr Biol 15:1235–1241, 2005. 30. Liu X, Bandyopadhyay BC, Singh BB, Groschner K, and Ambudkar IS. Molecular analysis of a store-operated and OAG sensitive non-selective cation channel: Heteromeric assembly of TRPC1-TRPC3. J Biol Chem, 2005. 31. Lockwich T, Singh BB, Liu X, and Ambudkar IS. Stabilization of cortical actin induces internalization of transient receptor potential 3 (Trp3)-associated caveolar Ca2+ signaling complex and loss of Ca2+ inﬂux without disruption of Trp3-inositol trisphosphate receptor association. J Biol Chem 276:42401–42408, 2001. 32. Lockwich TP, Liu X, Singh BB, Jadlowiec J, Weiland S, and Ambudkar IS. Assembly of Trp1 in a signaling complex associated with caveolin-scaffolding lipid raft domains. J Biol Chem 275:11934–11942, 2000. 33. Lutjens R, Igarashi M, Pellier V, Blasey H, Di Paolo G, Ruchti E, Pfulg C, Staple JK, Catsicas S, and Grenningloh G. Localization and targeting of SCG10 to the trans-Golgi apparatus and growth cone vesicles. Eur J Neurosci 12:2224–2234, 2000. 34. Ma HT, Patterson RL, van Rossum DB, Birnbaumer L, Mikoshiba K, and Gill DL. Requirement of the inositol trisphosphate receptor for activation of store-operated Ca2+ channels. Science 287:1647–1651, 2000. 35. Mehta D, Ahmmed GU, Paria BC, Holinstat M, Voyno-Yasenetskaya T, Tiruppathi C, Minshall RD, and Malik AB. RhoA interaction with inositol 1,4,5-trisphosphate receptor and transient receptor potential channel-1 regulates Ca2+ entry. Role in signaling increased endothelial permeability. J Biol Chem 278:33492–33500, 2003. 36. Mery L, Strauss B, Dufour JF, Krause KH, and Hoth M. The PDZ-interacting domain of TRPC4 controls its localization and surface expression in HEK293 cells. J Cell Sci 115:3497–3508, 2002. 37. Montell C. TRP channels in Drosophila photoreceptor cells. J Physiol 567:45–51, 2005. 38. Obukhov AG and Nowycky MC. TRPC5 activation kinetics are modulated by the scaffolding protein ezrin/radixin/moesin-binding phosphoprotein-50 (EBP50). J Cell Physiol 201:227–235, 2004. 39. Ordaz B, Tang J, Xiao R, Salgado A, Sampieri A, Zhu MX, and Vaca L. Calmodulin and calcium interplay in the modulation of TRPC5 channel activity. Identiﬁcation of a novel C-terminal domain for calcium/calmodulin-mediated facilitation. J Biol Chem 280:30788–30796, 2005. 40. Philipp S, Trost C, Warnat J, Rautmann J, Himmerkus N, Schroth G, Kretz O, Nastainczyk W, Cavalie A, Hoth M, and Flockerzi V. TRP4 (CCE1) protein is part of native calcium release-activated Ca2+-like channels in adrenal cells. J Biol Chem 275:23965–23972, 2000.

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41. Reading SA, Earley S, Waldron BJ, Welsh DG, and Brayden JE. TRPC3 mediates pyrimidine receptor-induced depolarization of cerebral arteries. Am J Physiol Heart Circ Physiol 288:H2055–2061, 2005. 42. Rosker C, Graziani A, Lukas M, Eder P, Zhu MX, Romanin C, and Groschner K. Ca2+ signaling by TRPC3 involves Na+ entry and local coupling to the Na+/Ca2+ exchanger. J Biol Chem, 2004. 43. Shieh BH and Zhu MY. Regulation of the TRP Ca2+ channel by INAD in Drosophila photoreceptors. Neuron 16:991–998, 1996. 44. Singh BB, Liu X, Tang J, Zhu MX, and Ambudkar IS. Calmodulin regulates Ca(2+)dependent feedback inhibition of store-operated Ca(2+) inﬂux by interaction with a site in the C-terminus of TrpC1. Mol Cell 9:739–750, 2002. 45. Singh BB, Lockwich TP, Bandyopadhyay BC, Liu X, Bollimuntha S, Brazer SC, Combs C, Das S, Leenders AG, Sheng ZH, Knepper MA, Ambudkar SV, and Ambudkar IS. VAMP2-dependent exocytosis regulates plasma membrane insertion of TRPC3 channels and contributes to agonist-stimulated Ca2+ inﬂux. Mol Cell 15:635–646, 2004. 46. Sinkins WG, Goel M, Estacion M, and Schilling WP. Association of immunophilins with mammalian TRPC channels. J Biol Chem 279:34521–34529, 2004. 47. Song X, Zhao Y, Narcisse L, Duffy H, Kress Y, Lee S, and Brosnan CF. Canonical transient receptor potential channel 4 (TRPC4) co-localizes with the scaffolding protein ZO-1 in human fetal astrocytes in culture. Glia 49:418–429, 2005. 48. Stamboulian S, Moutin MJ, Treves S, Pochon N, Grunwald D, Zorzato F, De Waard M, Ronjat M, and Arnoult C. Junctate, an inositol 1,4,5-trisphosphate receptor associated protein, is present in rodent sperm and binds TRPC2 and TRPC5 but not TRPC1 channels. Dev Biol 286:326–337, 2005. 49. Stein R, Mori N, Matthews K, Lo LC, and Anderson DJ. The NGF-inducible SCG10 mRNA encodes a novel membrane-bound protein present in growth cones and abundant in developing neurons. Neuron 1:463–476, 1988. 50. Strubing C, Krapivinsky G, Krapivinsky L, and Clapham DE. TRPC1 and TRPC5 form a novel cation channel in mammalian brain. Neuron 29:645–655, 2001. 51. Strubing C, Krapivinsky G, Krapivinsky L, and Clapham DE. Formation of novel TRPC channels by complex subunit interactions in embryonic brain. J Biol Chem 278:39014–39019, 2003. 52. Tang J, Lin Y, Zhang Z, Tikunova S, Birnbaumer L, and Zhu MX. Identiﬁcation of common binding sites for calmodulin and inositol 1,4,5-trisphosphate receptors on the carboxyl termini of trp channels. J Biol Chem 276:21303–21310, 2001. 53. Tang Y, Tang J, Chen Z, Trost C, Flockerzi V, Li M, Ramesh V, and Zhu MX. Association of mammalian trp4 and phospholipase C isozymes with a PDZ domaincontaining protein, NHERF. J Biol Chem 275:37559–37564, 2000. 54. Treves S, Franzini-Armstrong C, Moccagatta L, Arnoult C, Grasso C, Schrum A, Ducreux S, Zhu MX, Mikoshiba K, Girard T, Smida-Rezgui S, Ronjat M, and Zorzato F. Junctate is a key element in calcium entry induced by activation of InsP3 receptors and/or calcium store depletion. J Cell Biol 166:537–548, 2004. 55. Tsiokas L, Arnould T, Zhu C, Kim E, Walz G, and Sukhatme VP. Speciﬁc association of the gene product of PKD2 with the TRPC1 channel. Proc Natl Acad Sci USA 96:3934–3939, 1999. 56. Tsunoda S, Sun Y, Suzuki E, and Zuker C. Independent anchoring and assembly mechanisms of INAD signaling complexes in Drosophila photoreceptors. J Neurosci 21:150–158, 2001.

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57. van Rossum DB, Patterson RL, Sharma S, Barrow RK, Kornberg M, Gill DL, and Snyder SH. Phospholipase Cgamma1 controls surface expression of TRPC3 through an intermolecular PH domain. Nature 434:99–104, 2005. 58. Vazquez G, Lievremont JP, St JBG, and Putney JW, Jr. Human Trp3 forms both inositol trisphosphate receptor-dependent and receptor-independent store-operated cation channels in DT40 avian B lymphocytes. Proc Natl Acad Sci USA 98:11777–11782, 2001. 59. Vazquez G, Wedel BJ, Aziz O, Trebak M, and Putney JW, Jr. The mammalian TRPC cation channels. Biochim Biophys Acta 1742:21–36, 2004. 60. Voltz JW, Weinman EJ, and Shenolikar S. Expanding the role of NHERF, a PDZ-domain containing protein adapter, to growth regulation. Oncogene 20:6309–6314, 2001. 61. Wang T and Montell C. Rhodopsin formation in Drosophila is dependent on the PINTA retinoid-binding protein. J Neurosci 25:5187–5194, 2005. 62. Wedel BJ, Vazquez G, McKay RR, St JBG, and Putney JW, Jr. A calmodulin/inositol 1,4,5-trisphosphate (IP3) receptor-binding region targets TRPC3 to the plasma membrane in a calmodulin/IP3 receptor-independent process. J Biol Chem 278:25758– 25765, 2003. 63. Weinman EJ, Hall RA, Friedman PA, Liu-Chen LY, and Shenolikar S. The association of NHERF adaptor proteins with G protein-coupled receptors and receptor tyrosine kinases. Annu Rev Physiol, 2005. 64. Williams TM and Lisanti MP. The Caveolin genes: from cell biology to medicine. Ann Med 36:584–595, 2004. 65. Wu X, Babnigg G, and Villereal ML. Functional signiﬁcance of human trp1 and trp3 in store-operated Ca(2+) entry in HEK-293 cells. Am J Physiol Cell Physiol 278:C526–536, 2000. 66. Wu X, Babnigg G, Zagranichnaya T, and Villereal ML. The role of endogenous human Trp4 in regulating carbacol-induced calcium oscillations in HEK-293 cells. J Biol Chem 277:13597–13608, 2002. 67. Wu X, Zagranichnaya TK, Gurda GT, Eves EM, and Villereal ML. A TRPC1/TRPC3mediated increase in store-operated calcium entry is required for differentiation of H19-7 hippocampal neuronal cells. J Biol Chem 279:43392–43402, 2004. 68. Xiao B, Tu JC, Petralia RS, Yuan JP, Doan A, Breder CD, Ruggiero A, Lanahan AA, Wenthold RJ, and Worley PF. Homer regulates the association of group 1 metabotropic glutamate receptors with multivalent complexes of homer-related, synaptic proteins. Neuron 21:707–716, 1998. 69. Xu XZ, Chien F, Butler A, Salkoff L, and Montell C. TRPgamma, a drosophila TRPrelated subunit, forms a regulated cation channel with TRPL. Neuron 26:647–657, 2000. 70. Xu XZ, Li HS, Guggino WB, and Montell C. Coassembly of TRP and TRPL produces a distinct store-operated conductance. Cell 89:1155–1164, 1997. 71. Xu XZ, Moebius F, Gill DL, and Montell C. Regulation of melastatin, a TRP-related protein, through interaction with a cytoplasmic isoform. Proc Natl Acad Sci USA 98:10692–10697, 2001. 72. Yuan JP, Kiselyov K, Shin DM, Chen J, Shcheynikov N, Kang SH, Dehoff MH, Schwarz MK, Seeburg PH, Muallem S, and Worley PF. Homer binds TRPC family channels and is required for gating of TRPC1 by IP3 receptors. Cell 114:777–789, 2003. 73. Zagranichnaya TK, Wu X, and Villereal ML. Endogenous TRPC1, TRPC3, and TRPC7 proteins combine to form native store-operated channels in HEK-293 cells. J Biol Chem 280:29559–29569, 2005.

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74. Zhang SL, Yu Y, Roos J, Kozak JA, Deerinck TJ, Ellisman MH, Stauderman KA, and Cahalan MD. STIM1 is a Ca2+ sensor that activates CRAC channels and migrates from the Ca2+ store to the plasma membrane. Nature 437:902–905, 2005. 75. Zhang Z, Tang J, Tikunova S, Johnson JD, Chen Z, Qin N, Dietrich A, Stefani E, Birnbaumer L, and Zhu MX. Activation of Trp3 by inositol 1,4,5-trisphosphate receptors through displacement of inhibitory calmodulin from a common binding domain. Proc Natl Acad Sci USA 98:3168–3173, 2001. 76. Zhu MX. Multiple roles of calmodulin and other Ca2+-binding proteins in the functional regulation of TRP channels. Pﬂügers Arch, 2005.

25

Structural Insights into the Function of TRP Channels Rachelle Gaudet Harvard University

CONTENTS Structural Scaffold of TRP Channels....................................................................349 The Transmembrane Region..................................................................................350 The Pore Domain.......................................................................................350 The Sensor Domain ...................................................................................352 Cytosolic Domains ................................................................................................353 Ankyrin Repeats ........................................................................................353 MHR in TRPM Channels..........................................................................355 Enzymatic Domains...................................................................................355 The TRPML and TRPP Subfamilies.........................................................355 Oligomerization of Cytosolic Domains ................................................................356 Summary ................................................................................................................356 References..............................................................................................................356

STRUCTURAL SCAFFOLD OF TRP CHANNELS This chapter reviews our knowledge of the three-dimensional structure of TRP channels and how this information relates to channel function and regulation. Sequence analyses have revealed the domain composition of TRP channels. Because no structure has been determined for a complete TRP channel, I will ﬁrst break down TRP channels into component parts, or building blocks, and then discuss the structural information available for each, using the closest sequence homologues for which structures have been determined. TRP channels are part of the same channel superfamily as the voltage- and ligand-gated potassium channels.1 Because the highest sequence similarity between these and TRP channels is found in the transmembrane domain, all TRP channels are expected to form tetramers—homo- or heterotetramers—as functional units, like the voltage- and ligand-gated channels. Biochemical and biophysical analyses have conﬁrmed that several TRP channels are indeed tetrameric (e.g., see reference 2). The transmembrane domain of each TRP channel subunit is expected to contain six roughly membrane-spanning helical segments, S1 to S6—and therefore both the Nand C- terminal extensions are cytosolic. The transmembrane domain can be divided 349

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in two building blocks: the sensor, formed by helices S1–S4, and the pore, formed by helices S5 and S6. The pore forms a hole that spans the lipid membrane and passes ions and other hydrophilic molecules, to which the lipid membrane is otherwise impermeable. The pore contains a selectivity ﬁlter, the smallest constriction of pore, which dictates through its stereochemical and electrostatic properties what kind of molecules are allowed through the pore. The sensor perceives the signal(s) and transmits the information to the gate, the channel component that opens or closes the pore. The cytosolic domains of TRP channels contain regulatory components that can tune the channel opening propensity in a positive or negative fashion. Perhaps some of the most intriguing structural aspects of TRP channels are the diversity of their cytosolic domains. The TRP channel family is divided into seven subfamilies based on sequence similarity and function, and there is little homology in the N- and Cterminal cytosolic regions between subfamilies. Furthermore, while some common protein–protein interaction motifs can be identiﬁed in the cytosolic regions, such as ankyrin repeats, calmodulin-binding sites and PDZ domain–binding sites, other cytosolic segments appear novel by sequence analysis. The following sections highlight the current structural information on the TRP channel building blocks within the transmembrane region and the cytosolic domains.

THE TRANSMEMBRANE REGION THE PORE DOMAIN The putative S5 and S6 segments of TRP channels, upon tetramerization, are expected to form a central ion-conducting pore. The best characterized ion channel pore structure is that of the bacterial KcsA potassium channel.3–5 Other potassium channels more closely related to TRP channels, including the Kv1.2 Shaker channel, have a central pore structure very similar to that of KcsA.6–8 Therefore, these structures can be used as a model for the TRP channel pore. The KcsA structure has been described as an inverted teepee,3 with the S5 helices on the outside and inner S6 helices shaping the central pore opening and gate. The S6 helices splay out on the extracellular side of the membrane, leaving space for the S5–S6 linker sequence to shape the selectivity ﬁlter by forming a reentrant loop and short pore helix (Figure 25.1A and B). The KcsA selectivity ﬁlter sequence, TVGYGD, presents a set of oxygen groups precisely positioned to select for dehydrated K+ ions. Because the transmembrane topology of TRP channels and potassium channels is similar, it can be hypothesized that the S5–S6 linker also forms a reentrant loop shaping the selectivity ﬁlter in TRP channels, but that remains an open question requiring structure determination. In fact, the mammalian TRPV1–V4 channels have selectivity ﬁlter sequences similar to KcsA: all four channels have a TIG(M/L)GD sequence between their predicted S5 and S6 segments. However, KcsA is much more selective for potassium ions than these TRPV channels. For example, TRPV1 can pass Ca2+ > Mg2+ > Na+ K+ Cs+ and therefore has relatively high permeability for calcium ions (PCa/PNa = 9.60).9 How do these TRPV channels achieve the ability to pass diverse cations, both mono- and divalents? One possibility is that their scaffold of oxygens pointing into the pore is similar to

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FIGURE 25.1 (Color ﬁgure follows p. 234.) Structures of TRP channel-building blocks. The transmembrane domain of the Shaker channel (2A798), viewed from the intracellular side of the membrane (A) and from its side (B). The four subunits have distinct colors, and magenta spheres represent potassium ions in the selectivity ﬁlter. The S1–S4 helices form the sensor domain, connected to the S5–S6 pore domain through the S4–S5 linker. The S4–S5 linker forms a ring around the C-terminal end of the S6 helices and may trigger motion of the S6 helices to open or close the pore in response to voltage changes. Each sensor domain abuts the pore domain of the neighboring subunit, with their respective S4 and S5 helices in close contact. (C) Structure of the six ankyrin repeats in the N-terminal region of TRPV2 (2ETA44). Many TRP channels have ankyrin repeats in their N-terminal cytosolic regions. (D) Structure of the TRPM7 α-kinase domain, located at the C-terminal end of the channel (1IA951). An AMPPNP molecule is located in the active site, and a structural zinc ion is displayed as a grey sphere. (E) Structure of human NUDT9 with a ribose-5-phosphate molecule and two magnesium ions in the active site (1QVJ53). The C-terminal end of TRPM2 has 39 percent similarity to NUDT9.

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KcsA but is more ﬂexible, allowing the size of the cation to vary. Other TRP channels do not have a recognizable selectivity ﬁlter sequence homologous to potassium channels, and their linker between the predicted S5 and S6 helices in TRP channels varies greatly in length and sequence (reviewed in reference 10). Nonetheless, mutagenesis studies on TRPM4,11 TRPV5,12 and TRPV613 indicate that the residues in the S5–S6 linker and presumed selectivity ﬁlter are important determinants of the ion conductance or selectivity of these channels. The pore of potassium channels is opened by a kink in the S6 helices (Figure 25.1B8,14), and this mechanism is likely to be conserved in TRP channels. The TRP box, a short hydrophobic stretch conserved in the TRPC, TRPV, and TRPM subfamilies,15 is located just C-terminal of the putative S6 helix. Secondary structure prediction algorithms often predict that the S6 helix would extend beyond the membrane bilayer in the cytosolic side, including the TRP box. If these predictions are correct, the TRP box could, under some conditions, serve as a coiled-coil zipper that holds the channel in a closed conformation. Although the functional role of the TRP box is unclear, the TRP box regions of TRPM8, TRPV5, and TRPM5 have recently been implicated in sensing phosphatidylinositol 4,5-bisphosphate (PIP2) levels;16 a depletion of PIP2 caused the channels to close. Following the above structural hypothesis, PIP2 could sensitize channels by destabilizing the coiled coil, allowing other stimuli to stabilize an open channel conformation.

THE SENSOR DOMAIN The region encompassing putative transmembrane helices S1–S4 is named the sensor domain by analogy to the Shaker family of ion channels and other related voltagegated ion channels. The sensor domain of Shaker K+ channels contains multiple buried positively charged residues, which move within the membrane in response to changes in membrane polarization, sensing the voltage difference across the lipid membrane.17–19 These movements, in turn, are believed to open the gate inside the ion-conducting pore by splaying out the C-terminal and intracellular halves of the four S6 helices.14 Although there is less information about the location of various sensor modules in TRP channels, the name “sensor” also seems appropriate for the S1–S4 region in TRP channels. In fact, the capsaicin-binding site of TRPV1 has been mapped to the S2 and S3 segments,20 and the icilin-binding site of TRPM8 was mapped to the same region.21 Capsaicin and icilin induce the opening of the ion-conducting gate of TRPV1 and TRPM8, respectively. The sensor and pore regions of the potassium channels are clearly separate structural domains within the S1–S6 transmembrane region. This was made apparent in the structure of the bacterial KvAP potassium channel7 and more recently conﬁrmed in the structure of the Shaker channel.8 Applying the strict deﬁnition of a domain as an independently folding unit is certainly appropriate, because the sensor of KvAP could be expressed, puriﬁed, and crystallized as an isolated domain.7 The observation of two separate transmembrane domains within one protein actually represents a ﬁrst in membrane protein structural biology and therefore will likely cause a paradigm shift in the way we think about domain units in membrane proteins.

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How closely TRP channel sensor domains resemble that of the Shaker channel remains, again, an open question. But considering the sensor as an independent folding unit suggests that it will be possible to determine the structure of isolated sensors in the presence or absence of ligands to explore whether these ligands cause conformational changes within the isolated domain. In analogy to the current model of voltage gating in Shaker channels, it is also possible that the conformational change caused by stimuli consists rather of a rigid body movement of the sensor domain relative to the pore domain. It has also been reported that some TRP channels, speciﬁcally TRPV1, TRPV3, TRPM8, and TRPM4, are quite voltage sensitive, to the point where they could be described as voltage-gated channels with small gating charges22–24 (reviewed in reference 25). Voltage-gated potassium channels such as Shaker channels have four positively charged arginines in the S4 segment, which are known to sense transmembrane voltage.17,18 What are the corresponding voltage-sensitive residues in TRPV1, TRPV3, TRPM8, and TRPM4, and are they located in the sensor domain? TRPV1 does not have a readily identiﬁable “gating charge” residue in its sensor domain; the predicted S4 segment of TRPV channels does not contain any charged residue. Examining the other predicted transmembrane segments, an arginineglutamate pair is found in the S2 segment of TRPV1 (R474 and E478 in rat TRPV1). In TRPM channels, however, there are a number of positively and negatively charged residues in the S1–S4 region. Additional studies, including mutational analyses, are required to identify the gating charges in TRP channels and to determine whether they are localized in the S1–S4 region, which is the voltage-sensor domain in other voltage-gated ion channels of this topology.

CYTOSOLIC DOMAINS ANKYRIN REPEATS Ankyrin repeats are present in four subfamilies of TRP channels: the TRPA, TRPC, TRPN, and TRPV subfamilies. First identiﬁed in a pair of yeast transcription factors (Sw14 and Sw16), ankyrin repeat sequence motifs are found in over 3,600 proteins from all three kingdoms (bacteria, archea, and eukarya).26,27 The functions of ankyrin repeat proteins are diverse, ranging from transcription regulation and signaling to cytoskeleton assembly. The unifying characteristic of ankyrin repeats is that they mediate speciﬁc protein–protein interactions. In fact, no enzymatic function has been detected for any ankyrin repeat domain. The high-resolution structures of ankyrin repeat proteins reveal remarkable structural homology and regularity. The structure of each 33-residue ankyrin repeat sequence motif consists of two antiparallel α-helices followed by a long β-hairpin or “ﬁnger” loop projecting outward from the helices at a 90° angle. Consecutive repeats stack together to form a hydrophobic core that promotes protein folding and assembly, yielding an L-shaped elongated domain. The ankyrin repeat serves as a versatile scaffold for protein–protein interactions via nonconserved amino acids in the ﬁnger loops.28 No speciﬁc sequences or structural motifs, however, are universally recognized by ankyrin repeat proteins.

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The TRPA and TRPN channels, expressed in mechanosensitive hair cells, contain a large number of ankyrin repeats (up to 29 in Drosophila TRPN129), each of which is close to the ankyrin repeat consensus in sequence and in length. Why are there so many repeats in the TRPA and TRPN channels? A simple answer could be that there are multiple interacting protein targets. Another hypothesis is that a domain containing a large number of ankyrin repeats could behave as a physical spring.30 Molecular dynamics calculations have lent support to this hypothesis, showing that if a long ankyrin repeat domain is stretched in silico, the extension and stiffness of structures formed of 17 or 24 ankyrin repeats match those predicted for the gating spring in hair cells of the ear.31 This gating spring hypothesis, which elegantly explains the mechanosensitivity of hair cell conductance, can now be tested through further functional and structural studies of the TRPA and TRPN channels. In contrast to the TRPA and TRPN channels, the TRPC and TRPV channels contain a smaller number of repeats, four to six, each diverging signiﬁcantly from the ankyrin repeat sequence consensus deﬁned by analyses of ~4,000 repeats.26 The shorter ankyrin repeat domains of TRPC and TRPV family members may be protein– protein interaction domains mediating the assembly of signaling modules at the plasma membrane—either to relay signals sensed by the channels or to regulate their propensity to respond to signals. The ankyrin repeat domain of a TRPV channel is important to the integrity of the channel, as its deletion impairs assembly and trafﬁcking to the cell membrane.32–34 Similarly, deletion of the ankyrin repeats of TRPC1 impairs channel activity.35 Some candidate binding partners have been identiﬁed for TRPV ankyrin repeats. The TRPV1 N-terminus is important for calmodulin regulation of TRPV136 and associates in vitro and in vivo with synaptotagmin IX and snapin, two components of SNARE-dependent exocytosis in excitable cells.37 Two-hybrid screens have identiﬁed proteins that interact with the TRPV2 Nterminus, including the recombinase gene activator (RGA) and Acyl CoA binding domain protein 3 (ACBD3).38,39 Similarly, several proteins interact with the N-termini of TRPC channels, including a caveolin interaction with TRPC1,40 PLC-γ1 with TRPC3,41 and the dynamin superfamily member MxA with several TRPC channels.42 The ankyrin repeat domains of TRPV or TRPC channels could also interact intramolecularly with other channel regions, such as the C-terminal tail. Although there is no direct evidence of such an interaction, the expression of a TRPC2 splice variant consisting of the N-terminal cytosolic domain interferes with channel activity,43 suggesting that the N-terminal domain either interacts with some region of the full-length channel or displaces critical interaction partners. We and others have recently determined crystal structures of the ankyrin repeat domain (ARD) of the TRPV1 and TRPV2 ion channels (Figure 25.1C44,45). The TRPV1-ARD and TRPV2-ARD consist of six ankyrin repeat structural motifs, although only four of them are readily identiﬁable by sequence motif searches. There are several small sequence insertions that affect the overall ARD structure: ankyrin repeats one through three have unusually long and ﬂexible ﬁngers, whereas repeats ﬁve and six have unusually long outer helices. Furthermore, a large counterclockwise twist was observed in the stacking of repeats four and ﬁve. This twist breaks the regularity of the ankyrin repeat domain, altering the shape of surfaces available for

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interactions with proteins or other cellular ligands. Both solution studies and crystalpacking interactions indicate that the TRPV1- and TRPV2-ARDs do not form homooligomers,44 suggesting that the ARDs may be used for interactions with regulatory factors rather than in promoting tetrameric assembly of the ion channels. We predict that the TRPC channels will have similar ankyrin repeat domains of ﬁve or six repeats,44 although the structural details are likely to be different. The identiﬁcation of the true structural boundaries of ankyrin repeat domains of TRP channels is important to their functional analyses. Ankyrin repeat domains are known to have highly cooperative folding properties; mutation of a single residue within an individual repeat can affect the entire three-dimensional core structure.26 Short deletions within a TRP channel ankyrin repeat domain are therefore likely to be deleterious to the folding of the whole domain. In contrast, mutations in predicted ﬁnger-loop regions, which are the most common sites of protein–protein interactions for ankyrin repeats, could be very useful in determining binding surfaces and partners.

MHR

IN

TRPM CHANNELS

Unlike the TRPA, TRPC, TRPN, and TRPV channels, which contain ankyrin repeats in their N-terminal cytoplasmic domains, TRPM channels share four regions of high homology (TRPM homology regions—MHRs) in their N-terminal cytoplasmic segments. These MHRs are highly homologous within the TRPM family but have no recognizable sequence homology outside of the TRPM family. Little structural information can be inferred about these regions until their three-dimensional structures are determined. A S141L TRPM6 missense mutation in the MHR1 of TRPM6 causes hereditary hypomagnesemia46,47 and prevents oligomerization of the channel.48 Similarly, a splice variant of TRPM1 that contains only the N-terminal segment inhibits the translocation of full-length TRPM1 to the plasma membrane.49 The MHRs may therefore be involved in the oligomerization of the channels or in regulating transport to the plasma membrane.

ENZYMATIC DOMAINS Three TRP channels—TRPM2, TRPM6, and TRPM7—have C-terminal enzymatic domains (recently reviewed in reference 50). TRPM6 and TRPM7 both feature an α-kinase domain at their C-termini, and the structure of the TRPM7 α-kinase domain has been solved (Figure 25.1D51). The role of the kinase domain in the function of TRPM6 and TRPM7 channels and in their regulation by ATP is still a matter of debate.50 TRPM2 is regulated by ADP ribose through its C-terminal NUDT9 homology domain, which binds ADP ribose and has pyrophosphorylase activity.52 The structure of human NUDT9 has recently been solved (Figure 25.1E53), providing a scaffold for studying the function of the NUDT9 homology domain in TRPM2.53,54

THE TRPML

AND

TRPP SUBFAMILIES

The TRPP and TRPML channels form distant TRP channel subfamilies, with some homology to TRP channels in their transmembrane domain (recently reviewed in reference 55). Unlike most TRP channels, the TRPP and TRPML channels do not

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have very large cytosolic domains and have few recognizable structural motifs in these cytosolic regions. One unique structural feature of the TRPP and TRPML subfamilies is a large extracellular region linking the putative S1 and S2 transmembrane segments. In the TRPP subfamily, this extracellular region contains the “polycystin” motif, although no function has yet been attributed to this conserved region.

OLIGOMERIZATION OF CYTOSOLIC DOMAINS The structures of cytosolic domains of voltage- and ligand-gated channels have demonstrated that many of these cytosolic domains form oligomers. Most form tetramers with fourfold symmetry, mirroring the tetrameric transmembrane domains. Examples include the T1 domain of Shaker channels,56 the Ca2+-binding domain of Ca2+-gated channels,14 the N- and C-terminal regions of G-protein-coupled inward rectifying potassium channels,57 and the C-terminal cyclic nucleotide-binding domain of a mammalian cyclic nucleotide-gated channel.58 However, some form dimers instead: the cyclic nucleotide-binding domain of the bacterial cyclic nucleotidegated channel MlotiK1 is an example of a dimeric domain of a tetrameric potassium channel.59 In all cases, the oligomerization is thought important for the regulation of the channel, by enabling cooperativity between the subunits through concerted conformational changes. There are several indications that cytosolic domains of TRP channels may also be oligomers. As stated above, the MHR region of TRPM channels and the Nterminal region of TRPC2 are important for the assembly of the respective channels, suggesting that the regions may themselves tetramerize. Furthermore, the ankyrin repeats of TRPV5 and TRPV6 have also been implicated in oligomerization of the channels.32,33 Finally, the C-terminal domain of TRPV1 may also oligomerize.60 In most cases, however, the evidence for—or suggestion of—oligomerization of the cytosolic domains of TRP channels is still only indirect. Studying the oligomerization properties of cytosolic domains can therefore provide important insights into the structure and function of TRP channels and the roles of their incredibly diverse cytosolic domains.

SUMMARY The structures of TRP channel building blocks or their homologues are proving useful in deciphering the inner workings of TRP channels. However, there is still much to be learned about how these building blocks work together to execute the various physiological functions of this fascinating family of ion channels. This will require the structure determination of complete TRP channels in different conformations representing distinct functional states.

REFERENCES 1. Harteneck, C., Plant, T.D., and Schultz, G., From worm to man: three subfamilies of TRP channels, Trends Neurosci. 23 (4), 159–66, 2000.

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2. Kedei, N., Szabo, T., Lile, J.D., Treanor, J.J., Olah, Z., Iadarola, M.J., and Blumberg, P.M., Analysis of the native quaternary structure of vanilloid receptor 1, J. Biol. Chem. 276 (30), 28613–19, 2001. 3. Doyle, D.A., Morais–Cabral, J., Pfuetzner, R.A., Kuo, A., Gulbis, J.M., Cohen, S.L., Chait, B.T., and MacKinnon, R., The structure of the potassium channel: molecular basis of K+ conduction and selectivity, Science 280 (5360), 69–77, 1998. 4. Zhou, Y., Morais-Cabral, J.H., Kaufman, A., and MacKinnon, R., Chemistry of ion coordination and hydration revealed by a K+ channel—Fab complex at 2.0 Å resolution, Nature 414 (6859), 43–48, 2001. 5. Morais-Cabral, J.H., Zhou, Y., and MacKinnon, R., Energetic optimization of ion conduction rate by the K+ selectivity ﬁlter, Nature 414 (6859), 37–42, 2001. 6. Jiang, Y., Lee, A., Chen, J., Cadene, M., Chait, B.T., and MacKinnon, R., Crystal structure and mechanism of a calcium-gated potassium channel, Nature 417 (6888), 515–22, 2002. 7. Jiang, Y., Lee, A., Chen, J., Ruta, V., Cadene, M., Chait, B.T., and MacKinnon, R., X-ray structure of a voltage-dependent K+ channel, Nature 423 (6935), 33–41, 2003. 8. Long, S.B., Campbell, E.B., and Mackinnon, R., Crystal structure of a mammalian voltage-dependent Shaker family K+ channel, Science 309 (5736), 897–903, 2005. 9. Caterina, M.J., Schumacher, M.A., Tominaga, M., Rosen, T.A., Levine, J.D., and Julius, D., The capsaicin receptor: a heat-activated ion channel in the pain pathway, Nature 389 (6653), 816–24, 1997. 10. Voets, T. and Nilius, B., The pore of TRP channels: trivial or neglected? Cell Calcium 33 (5–6), 299–302, 2003. 11. Nilius, B., Prenen, J., Janssens, A., Owsianik, G., Wang, C., Zhu, M.X., and Voets, T., The selectivity ﬁlter of the cation channel TRPM4, J. Biol. Chem. 280 (24), 22899–906, 2005. 12. Dodier, Y., Banderali, U., Klein, H., Topalak, O., Daﬁ, O., Simoes, M., Bernatchez, G., Sauve, R., and Parent, L., Outer pore topology of the ECaC–TRPV5 channel by cysteine scan mutagenesis, J. Biol. Chem. 279 (8), 6853–62, 2004. 13. Voets, T., Janssens, A., Droogmans, G., and Nilius, B., Outer pore architecture of a Ca2+-selective TRP channel, J. Biol. Chem. 279 (15), 15223–30, 2004. 14. Jiang, Y., Lee, A., Chen, J., Cadene, M., Chait, B.T., and MacKinnon, R., The open pore conformation of potassium channels, Nature 417 (6888), 523–26, 2002. 15. Montell, C., Physiology, phylogeny, and functions of the TRP superfamily of cation channels, Science's STKE, 2001. 16. Rohacs, T., Lopes, C.M., Michailidis, I., and Logothetis, D.E., PI(4,5)P2 regulates the activation and desensitization of TRPM8 channels through the TRP domain, Nat. Neurosci. 8 (5), 626–34, 2005. 17. Aggarwal, S.K. and MacKinnon, R., Contribution of the S4 segment to gating charge in the Shaker K+ channel, Neuron 16 (6), 1169–77, 1996. 18. Seoh, S.A., Sigg, D., Papazian, D.M., and Bezanilla, F., Voltage-sensing residues in the S2 and S4 segments of the Shaker K+ channel, Neuron 16 (6), 1159–67, 1996. 19. Tombola, F., Pathak, M.M., and Isacoff, E.Y., How far will you go to sense voltage? Neuron 48 (5), 719–25, 2005. 20. Jordt, S.E. and Julius, D., Molecular basis for species-speciﬁc sensitivity to ‘‘hot’’ chili peppers, Cell 108 (3), 421–30, 2002. 21. Chuang, H.-h., Neuhausser, W.M., and Julius, D., The super-cooling agent icilin reveals a mechanism of coincidence detection by a temperature-sensitive TRP channel, Neuron 43 (6), 859–69, 2004.

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22. Voets, T., Droogmans, G., Wissenbach, U., Janssens, A., Flockerzi, V., and Nilius, B., The principle of temperature-dependent gating in cold- and heat-sensitive TRP channels, Nature 430 (7001), 748–54, 2004. 23. Chung, M.K., Guler, A.D., and Caterina, M.J., Biphasic currents evoked by chemical or thermal activation of the heat-gated ion channel, TRPV3, J. Biol. Chem. 280 (16), 15928–41, 2005. 24. Nilius, B., Prenen, J., Droogmans, G., Voets, T., Vennekens, R., Freichel, M., Wissenbach, U., and Flockerzi, V., Voltage dependence of the Ca2+-activated cation channel TRPM4, J. Biol. Chem. 278 (33), 30813–20, 2003. 25. Nilius, B., Talavera, K., Owsianik, G., Prenen, J., Droogmans, G., and Voets, T., Gating of TRP channels: a voltage connection? J. Physiol. 567 (Pt. 1), 35–44, 2005. 26. Mosavi, L.K., Minor, D.L., Jr., and Peng, Z.Y., Consensus-derived structural determinants of the ankyrin repeat motif, Proc. Natl. Acad. Sci. USA 99 (25), 16029–34, 2002. 27. Schultz, J., Copley, R.R., Doerks, T., Ponting, C.P., and Bork, P., SMART: a webbased tool for the study of genetically mobile domains, Nucleic Acids Res. 28 (1), 231–34, 2000. 28. Kohl, A., Binz, H.K., Forrer, P., Stumpp, M.T., Pluckthun, A., and Grutter, M.G., Designed to be stable: crystal structure of a consensus ankyrin repeat protein, Proc. Natl. Acad. Sci. USA 100 (4), 1700–1705, 2003. 29. Walker, R.G., Willingham, A.T., and Zuker, C.S., A Drosophila mechanosensory transduction channel, Science 287 (5461), 2229–34, 2000. 30. Howard, J. and Bechstedt, S., Hypothesis: a helix of ankyrin repeats of the NOMPCTRP ion channel is the gating spring of mechanoreceptors, Curr. Biol. 14 (6), R224–26, 2004. 31. Sotomayor, M., Corey, D.P., and Schulten, K., In search of the hair-cell gating spring elastic properties of ankyrin and cadherin repeats, Structure (Camb) 13 (4), 669–82, 2005. 32. Chang, Q., Gyftogianni, E., van de Graaf, S.F., Hoefs, S., Weidema, F.A., Bindels, R.J., and Hoenderop, J.G., Molecular determinants in TRPV5 channel assembly, J. Biol. Chem. 279 (52), 54304–11, 2004. 33. Erler, I., Hirnet, D., Wissenbach, U., Flockerzi, V., and Niemeyer, B.A., Ca2+-selective transient receptor potential V channel architecture and function require a speciﬁc ankyrin repeat, J. Biol. Chem. 279 (33), 34456–63, 2004. 34. Jung, J., Lee, S.Y., Hwang, S.W., Cho, H., Shin, J., Kang, Y.S., Kim, S., and Oh, U., Agonist recognition sites in the cytosolic tails of vanilloid receptor 1, J. Biol. Chem. 277 (46), 44448–54, 2002. 35. Engelke, M., Friedrich, O., Budde, P., Schafer, C., Niemann, U., Zitt, C., Jungling, E., Rocks, O., Luckhoff, A., and Frey, J., Structural domains required for channel function of the mouse transient receptor potential protein homologue TRP1beta, FEBS Lett. 523 (1–3), 193–99, 2002. 36. Rosenbaum, T., Gordon-Shaag, A., Munari, M., and Gordon, S.E., Ca2+/calmodulin modulates TRPV1 activation by capsaicin, J. Gen. Physiol. 123 (1), 53–62, 2004. 37. Morenilla-Palao, C., Planells-Cases, R., Garcia-Sanz, N., and Ferrer-Montiel, A., Regulated exocytosis contributes to protein kinase C potentiation of vanilloid receptor activity, J. Biol. Chem. 279 (24), 25665–72, 2004. 38. Barnhill, J.C., Stokes, A.J., Koblan-Huberson, M., Shimoda, L.M., Muraguchi, A., Adra, C.N., and Turner, H., RGA protein associates with a TRPV ion channel during biosynthesis and trafﬁcking, J. Cell Biochem. 91 (4), 808–20, 2004.

Structural Insights into the Function of TRP Channels

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39. Stokes, A.J., Wakano, C., Del Carmen, K.A., Koblan-Huberson, M., and Turner, H., Formation of a physiological complex between TRPV2 and RGA protein promotes cell surface expression of TRPV2, J. Cell Biochem. 94 (4), 669–83, 2005. 40. Brazer, S.C., Singh, B.B., Liu, X., Swaim, W., and Ambudkar, I.S., Caveolin-1 contributes to assembly of store-operated Ca2+ inﬂux channels by regulating plasma membrane localization of TRPC1, J. Biol. Chem. 278 (29), 27208–15, 2003. 41. van Rossum, D.B., Patterson, R.L., Sharma, S., Barrow, R.K., Kornberg, M., Gill, D.L., and Snyder, S.H., Phospholipase C[gamma]1 controls surface expression of TRPC3 through an intermolecular PH domain, Nature 434 (7029), 99–104, 2005. 42. Lussier, M.P., Cayouette, S., Lepage, P.K., Bernier, C.L., Francoeur, N., St-Hilaire, M., Pinard, M., and Boulay, G., MxA, a member of the dynamin superfamily, interacts with the ankyrin-like repeat domain of TRPC, J. Biol. Chem. 280 (19), 19393–19400, 2005. 43. Chu, X., Tong, Q., Wozney, J., Zhang, W., Cheung, J.Y., Conrad, K., Mazack, V., Stahl, R., Barber, D.L., and Miller, B.A., Identiﬁcation of an N-terminal TRPC2 splice variant which inhibits calcium inﬂux, Cell Calcium 37 (2), 173–82, 2005. 44. Jin, X., Touhey, J., and Gaudet, R., Structure of the N-terminal ankyrin repeat domain of the TRPV2 ion channel, J. Biol. Chem, in press, 2006. 45. McCleverty, C., Koesema, E., Patapoutian, A., Lesley, S.A., and Kreusch, A., Crystal structure of the human TRPV2 channel ankyrin repeat domain, Protein Science, in press, 2006. 46. Walder, R.Y., Landau, D., Meyer, P., Shalev, H., Tsolia, M., Borochowitz, Z., Boettger, M.B., Beck, G.E., Englehardt, R.K., Carmi, R., and Shefﬁeld, V.C., Mutation of TRPM6 causes familial hypomagnesemia with secondary hypocalcemia, Nat. Genet. 31 (2), 171–74, 2002. 47. Schlingmann, K.P., Weber, S., Peters, M., Niemann Nejsum, L., Vitzthum, H., Klingel, K., Kratz, M., Haddad, E., Ristoff, E., Dinour, D., Syrrou, M., Nielsen, S., Sassen, M., Waldegger, S., Seyberth, H.W., and Konrad, M., Hypomagnesemia with secondary hypocalcemia is caused by mutations in TRPM6, a new member of the TRPM gene family, Nat. Genet. 31 (2), 166–70, 2002. 48. Chubanov, V., Waldegger, S., Mederos y Schnitzler, M., Vitzthum, H., Sassen, M.C., Seyberth, H.W., Konrad, M., and Gudermann, T., Disruption of TRPM6/TRPM7 complex formation by a mutation in the TRPM6 gene causes hypomagnesemia with secondary hypocalcemia, Proc. Natl. Acad. Sci. USA 101 (9), 2894–99, 2004. 49. Xu, X.Z., Moebius, F., Gill, D.L., and Montell, C., Regulation of melastatin, a TRPrelated protein, through interaction with a cytoplasmic isoform, Proc. Natl. Acad. Sci. USA 98 (19), 10692–97, 2001. 50. Scharenberg, A., TRPM2 and TRPM7: channel/enzyme fusions to generate novel intracellular sensors, Pﬂügers Arch. 451 (1), 220–27, 2005. 51. Yamaguchi, H., Matsushita, M., Nairn, A.C., and Kuriyan, J., Crystal structure of the atypical protein kinase domain of a TRP channel with phosphotransferase activity, Mol. Cell 7 (5), 1047–57, 2001. 52. Perraud, A.L., Fleig, A., Dunn, C.A., Bagley, L.A., Launay, P., Schmitz, C., Stokes, A.J., Zhu, Q., Bessman, M.J., Penner, R., Kinet, J.P., and Scharenberg, A.M., ADPribose gating of the calcium-permeable LTRPC2 channel revealed by Nudix motif homology, Nature 411 (6837), 595–99, 2001. 53. Shen, B.W., Perraud, A.L., Scharenberg, A., and Stoddard, B.L., The crystal structure and mutational analysis of human NUDT9, J. Mol. Biol. 332 (2), 385–98, 2003. 54. Kolisek, M., Beck, A., Fleig, A., and Penner, R., Cyclic ADP-ribose and hydrogen peroxide synergize with ADP-ribose in the activation of TRPM2 channels, Mol. Cell 18 (1), 61–69, 2005.

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55. Qian, F. and Noben-Trauth, K., Cellular and molecular function of mucolipins (TRPML) and polycystin 2 (TRPP2), Pﬂügers Arch 451 (1), 277–85, 2005. 56. Kreusch, A., Pfafﬁnger, P.J., Stevens, C.F., and Choe, S., Crystal structure of the tetramerization domain of the Shaker potassium channel, Nature 392 (6679), 945–48, 1998. 57. Nishida, M. and MacKinnon, R., Structural basis of inward rectiﬁcation: cytoplasmic pore of the G protein-gated inward rectiﬁer GIRK1 at 1.8 A resolution, Cell 111 (7), 957–65, 2002. 58. Zagotta, W.N., Olivier, N.B., Black, K.D., Young, E.C., Olson, R., and Gouaux, E., Structural basis for modulation and agonist speciﬁcity of HCN pacemaker channels, Nature 425 (6954), 200–205, 2003. 59. Clayton, G.M., Silverman, W.R., Heginbotham, L., and Morais-Cabral, J.H., Structural basis of ligand activation in a cyclic nucleotide-regulated potassium channel, Cell 119 (5), 615–27, 2004. 60. Garcia-Sanz, N., Fernandez-Carvajal, A., Morenilla-Palao, C., Planells-Cases, R., Fajardo-Sanchez, E., Fernandez-Ballester, G., and Ferrer-Montiel, A., Identiﬁcation of a tetramerization domain in the C-terminus of the vanilloid receptor, J. Neurosci. 24 (23), 5307–14, 2004.

26

Functional Signiﬁcance of Transient Receptor Potential Channels in Vascular Function Scott Earley* Stacey Reading Joseph E. Brayden University of Vermont College of Medicine

CONTENTS Introduction............................................................................................................362 TRP Channel Function in Vascular Smooth Muscle Cells...................................363 Vasoconstrictor Agonist-Induced TRP Channel Activation in Smooth Muscle......................................................................................363 Vasodilator Response to EETs Mediated by TRP Channel Activation in Smooth Muscle .....................................................366 Pressure-Induced TRP Channel Activation in Vascular Smooth Muscle ..........................................................................................366 Store-Operated TRP Channel Activation in Vascular Smooth Muscle ..........................................................................................368 TRP Channel Function in Vascular Endothelial Cells..........................................369 Presence of TRP Channels in the Endothelium........................................369 TRP Channels and Endothelial Cell Calcium Entry ................................369 TRP Channels in the Proliferative Response of the Endothelium to Hypoxia.................................................................371 TRP Channels and Regulation of Capillary Permeability........................371 Role of TRP Channels in Endothelial Cell Response to Temperature Change..............................................................................371 TRP Channels as Sensors of Endothelial Oxidative Stress......................372 Summary and Future Perspectives ........................................................................372 References..............................................................................................................372 * Present afﬁliation: Colorado State University, Fort Collins.

361

362

TRP Ion Channel Function in Sensory Transduction

INTRODUCTION Membrane ion channels are critical mediators of vascular endothelial and smooth muscle cell function and regulate diverse properties of blood vessels such as arterial tone, angiogenesis, and permeabilty. Although physiological roles for many ion channels and transporters are well established, the functional signiﬁcance of transient receptor potential (TRP) channels in the vasculature is just beginning to be elucidated. This chapter summarizes the current understanding of these cation channels in vascular function. For a more in-depth overview of TRP channel biology, the reader is directed to several excellent general reviews covering this topic.1–4 As shown in Table 26.1, the mammalian TRP superfamily of cation channels contains at least 22 genes grouped into three major subfamilies based on sequence

TABLE 26.1 Summary of TRP Channels in Vascular Tissue Channel

Present in Arteries?

TRPC1

yes

TRPC2

yes

TRPC3

yes

TRPC4

yes

TRPC5

yes

TRPC6

yes

TRPC7

yes

TRPV1 TRPV2

yes yes

TRPV3 TRPV4

yes yes

TRPV5 TRPV6 TRPM1 TRPM2 TRPM3 TRPM4 TRPM5 TRPM6 TRPM7 TRPM8

no no yes yes yes yes yes yes yes yes

Cell Type

Function?

smooth muscle48 endothelium56 smooth muscle23 endothelium23 smooth muscle9 endothelium61 smooth muscle9 endothelium55,56,70 smooth muscle71 endothelium56,70 smooth muscle28 endothelium56,70 smooth muscle9 endothelium70 ? smooth muscle42 endothelium53 ? smooth muscle36 endothelium53,54 — — ? ? ? smooth muscle44,45 ? ? ? ?

SOC48 ? ? UTP-induced depolarization31 SOC59 SOC51,61 endothelial permeability62 ? PE-induced constriction myogenic tone 33 ?

28

? stretch-induced cation current42 ? EETs receptor36,65 thermosensitivity68 — — ? ? ? myogenic tone45 ? ? ? ?

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363

homology: TRPV (vanilloid), TRPC (canonical), and TRPM (melastatin). Three additional subfamilies (the “distant TRPs”), TRPP (polycystin), TRPML (mucolipin), and TRPA have been proposed, bringing the total number of TRPrelated proteins to around 30. Although these channels were initially described in sensory neurons, it is now thought that most cell types express several TRP genes. TRP proteins are expressed as six transmembrane-domain polypeptide subunits, and it is believed that four subunits assemble in the plasma membrane to form functional channels. All TRP channels are cation permeable, and most are not selective for monovalent versus divalent ions. Exceptions include TRPV5 and TRPV6, which display signiﬁcant speciﬁcity for Ca2+ ions, and TRPM4 and TRPM5, which are highly selective for monovalent cations and impermeant to Ca2+. TRP channels are activated by a variety of stimuli, including changes in pressure, temperature, osmolarity, and intracellular Ca2+. Fatty acids and receptor-dependent vasoconstrictor agonists also activate vascular TRP channels. This diversity of ionic conductivity and activating mechanisms is consistent with the possibility that members of the TRP superfamily may contribute to regulation of a variety of physiological systems. Elucidation of functional roles for TRP channels in vascular cells may be hindered by the complex molecular biology of the superfamily. Biophysical properties of TRP channels have been investigated in patch-clamp experiments employing cultured cells expressing cloned TRP subunit genes. Under these conditions, most functional channels assemble from four identical TRP subunits. However, when multiple TRP subunits are coexpressed, assembly of tetramers composed of two or more TRP subunit proteins can form heteromeric channels5,6 with novel properties.7,8 Because most cells express multiple TRP subunits, it is likely that heteromeric channels exist in vivo. Furthermore, splice variants of TRP mRNAs in smooth muscle have been reported,9 potentially increasing the number of individual subunits available for coassembly. Because TRP molecular variety could result in an assortment of channels with a wide array of functional properties in native cells, unraveling the physiological consequences of TRP channel diversity presents a major challenge. In the following paragraphs, evidence for the presence and possible functional roles of TRP channels in vascular smooth muscle and endothelial cells are discussed.

TRP CHANNEL FUNCTION IN VASCULAR SMOOTH MUSCLE CELLS VASOCONSTRICTOR AGONIST-INDUCED TRP CHANNEL ACTIVATION IN SMOOTH MUSCLE Vascular smooth muscle cells contract when exposed to a variety of excitatory neurotransmitters and hormones such as endothelin, histamine, norepinephrine, serotonin, and purine and pyrimidine nucleotides. Many of these constrictor agonists bind to G-protein- or tyrosine kinase–coupled receptors that activate phospholipase C (PLC) to initiate the conversion of phosphatidylinositol 4,5-bisphosphate into inositol-1,4,5-trisphosphate (IP3) and diacylglycerol (DAG) at the sarcolemmal membrane.10 PLC-coupled agonists raise cytosolic Ca2+ ([Ca2+]c) with a characteristic early transient phase followed by a secondary sustained phase.

364

TRP Ion Channel Function in Sensory Transduction

IP3-mediated Ca2+ release from the sarcoplasmic reticulum contributes to the early large increase in [Ca2+]c.11,12 Ca2+ inﬂux across the sarcolemmal membrane contributes to the maintenance of cytosolic Ca2+ during the sustained phase as evidenced by the fact that elevated [Ca2+]c and vasoconstriction are not maintained in the absence of extracellular Ca2+.13 Extracellular Ca2+ inﬂux occurs in part through well-deﬁned voltage-gated Ca2+ channels14 as well as through channels gated independently of the membrane potential.15 These nonselective cation channels coupled to PLC, possibly through DAG, appear to play a role in the extracellular Ca2+ inﬂux by directly permitting Ca2+ inﬂux or by permitting the inﬂux of monovalent cations that depolarize the smooth muscle cell.16–19 The molecular identities of these channels are not yet known. Accumulating evidence supports the idea that some of these channels may in fact be mammalian homologues of TRP channels ﬁrst identiﬁed in photoreceptors of Drosophila melanogaster. In Drosophila, light causes a sustained depolarization of photoreceptors through a signal transduction pathway mediated by PLC.20 Two conductances—one that is Ca2+ selective and another that is permeable to both Ca2+ and Na+—are responsible for light-induced depolarization. Gene products encoded by trp and trpl (TRP-like) are believed to be responsible for these conductances by forming membrane channels that have structural homologues with voltage-gated channels.21 Of the three identiﬁed mammalian TRP channel families, members of the TRPC family share the greatest similarity to Drosophila TRP and TRPL channels.4 Thus, TRPC channels may be responsible for voltage-independent extracellular Ca2+ inﬂux in vascular smooth muscle exposed to PLC-coupled agonists. There is evidence to show that most TRPC channels (TRPC2 is the exception) are activated by stimulation of PLC, formation of DAG, or exposure to DAG analogues. TRPC channels appear to be widespread in the mammalian vasculature and mRNA, for all seven TRPC channels have been detected in venous and arterial smooth muscle from a variety of vascular beds.3,9,22,23 TRPC1, TRPC3, and TRPC6 are abundant in vascular smooth muscle, and these proteins have been detected by Western blot in every vascular bed so far examined. TRPC channels are divided into four subfamilies based on amino acid homology: TRPC1, TRPC2, TRPC4/5 (60 percent homology), and TRPC3/6/7 (~80 percent homology).24 Functional channels form as tetramers of individual TRPC proteins. All TRPC subunits can combine to form homomeric channels,25 and heteromeric channels can result when TRPC proteins combine with other members of their own subfamily6 or when TRPC1 combines with either TRPC4/56,8 or TRPC3.26,27 The combinatorial rules governing the assembly of TRPC proteins into functional channels in vascular smooth muscle cells are not known. It is likely that TRPC proteins assemble to form both homomeric and heteromeric channels. Elucidation of TRPC channel function in native vascular cells has been impeded by the complex molecular biology of the TRP superfamily and by the lack of selective pharmacological inhibitors. A few studies using genetic methods to downregulate TRPC expression in intact vascular tissue have yielded valuable information. For example, Inoue et al.28 observed that exposure to phenylephrine (PE, a vasoconstrictor agonist that activates α1-adrenoreceptors coupled to PLC) produced a current in vascular smooth muscle cells isolated from the rabbit portal vein that displayed dual

Functional Signiﬁcance of Transient Receptor Potential Channels

365

rectiﬁcation; the channel was permeable to divalent cations and had a unitary conductance of 25 to 30 pS. This channel could also be activated by DAG, and the channel activity was potentiated by ﬂufenamte (a nonspeciﬁc cation channel inhibitor that “uniquely” enhances α1-adrenoreceptor nonselective cation currents in the rabbit portal vein) and by extracellular Ca2+. The channel was inhibited by other nonselective cation channel inhibitors including Cd2+, La3+, Gd3+, SK&F96365, and amiloride. PEinduced currents with virtually identical properties to those found in the native cells were recorded from HEK cells overexpressing cloned TRPC6. Downregulation of TRPC6 expression in primary cultured portal vein myocytes using antisense oligodeoxynucleotides resulted in suppression of α1-adrenoreceptor-activated cation currents. These ﬁndings show that TRPC6 channels are critical mediators of PE-induced responses in portal vein smooth muscle cells. A current with biophysical and pharmacological properties similar to that described by Inoue et al.28 was recorded in the smooth muscle–derived A7r5 cell line exposed to vasopressin or membranepermeable analogues of DAG.29 Northern blots of cDNA probes for all seven TRPCs revealed that only TRPC1 and TRPC6 were present in these cells. Because the current showed properties characteristic of TRPC6, and because there is no evidence to support the presence of a TRPC1/6 heteromeric channel, it is likely that the agonistinduced current was mediated by TRPC6 channels. Uridine trisphosphate (UTP) causes depolarization and contraction of both coronary and cerebrovascular smooth muscle cells.30,31 We found that antisense oligonucleotide suppression of TRPC3 expression in cerebral arteries resulted in a diminished UTP-induced depolarization and vasoconstriction.31 In addition, UTP-activated whole-cell cation currents were greatly suppressed in cerebral artery myocytes following antisense-mediated TRPC3 downregulation. Furthermore, although about 50 percent of the UTP-induced, TRPC3-mediated constriction appears to be due to the depolarization and activation of L-type calcium channels, our recent observations suggest that much of the balance of the vasoconstriction is due to direct calcium entry via the TRPC3 channel per se32 (see Figure 26.1). Findings from our laboratory also show that, in rat cerebral arteries, TRPC3 and TRPC6 are coupled to distinct excitatory stimuli. TRPC3 (and not TRPC6) is involved in UTP-mediated depolarization and vasoconstriction, whereas TRPC6 (and not TRPC3) is involved in pressure-induced depolarization and vasoconstriction.31,33

FIGURE 26.1 TRP channels that are receptor coupled and that directly or indirectly mediate calcium entry in vascular smooth muscle cells.

366

TRP Ion Channel Function in Sensory Transduction

These ﬁndings suggest that individual TRPC channel isoforms may be activated by different stimuli (i.e., agonist versus pressure) in the same vascular bed. In addition to being involved in normal vascular smooth muscle cell function, a few studies have shown that TRPC expression is altered in vascular smooth muscle during pathological conditions or during environmental stress. For example, Yu et al.22 showed that TRPC3 and TRPC6 expression was signiﬁcantly greater in the pulmonary artery smooth muscle cells of patients with idiopathic pulmonary hypertension when compared with healthy individuals or patients with secondary pulmonary hypertension. Other investigators reported that arteries undergoing organ culture and balloon dilation showed increased TRPC1 and TRPC6 expression, whereas TRPC3 expression was decreased.34 Another study demonstrated that chronic hypoxia increased TRPC1 and TRPC6 expression in intralobar artery smooth muscle cells in rabbits.35 The implication of this observed plasticity in TRPC expression is that TRPC channels are involved in normal vascular smooth muscle function and may also play a role in vascular dysfunction and pathology. Future exploration of agonist-TRPC channel interaction will likely yield exciting new information on vascular smooth muscle cell function as well as insight into a variety of vascular pathologies.

VASODILATOR RESPONSE TO EETS MEDIATED ACTIVATION IN SMOOTH MUSCLE

BY

TRP CHANNEL

TRPV4 is present in arterial cells. A recent report by Earley et al.36 indicates that TRPV4 is involved in vasodilation induced by epoxyeicosatrienoic acid compounds (EETs). These vasodilatory factors are produced by the vascular endothelium and have been proposed to be one form of endothelium-derived hyperpolarizing factor. Earley et al. found that EET-induced vasodilation of cerebral arteries involves a cascade of responses beginning with the activation of TRPV4 channels located in the smooth muscle cell membrane. Calcium entry via the activated TRPV4 channels then appears to activate the release of calcium from the sarco-endoplasmic reticulum in the form of local calcium release events termed calcium sparks. EET-generated Ca2+ sparks then activate nearby sarcolemmal large conductance Ca2+-sensitive K+ (BKCa) channels and increase the frequency of transient K+ currents (referred to as “spontaneous transient outward currents” or STOCs). The EET-induced increases in STOC frequency results in smooth muscle hyperpolarization and vasodilation. The conclusion of this work is that TRPV4 forms a novel Ca2+-signaling complex with ryanodine receptors and BKCa channels that elicits smooth muscle hyperpolarization and arterial dilation via Ca2+-induced Ca2+ release in response to an endothelialderived factor (see Figure 26.1).

PRESSURE-INDUCED TRP CHANNEL ACTIVATION SMOOTH MUSCLE

IN

VASCULAR

Resistance arteries constrict in response to increasing intraluminal pressure.37 This phenomenon, known as the vascular myogenic response, is an important autoregulatory mechanism that allows blood ﬂow to remain relatively constant despite large

Functional Signiﬁcance of Transient Receptor Potential Channels

367

changes in intravascular pressure.38 Myogenic constriction results from activation of voltage-dependent Ca2+ channels39 secondary to smooth muscle cell membrane depolarization.40 Activation of mechanosensitive ion channels appears to play a central role in pressure-induced depolarization and myogenic constriction. Consistent with this hypothesis, a number of early reports describe Gd3+-sensitive stretchactivated nonselective cation channels in smooth muscle cells. These stretchactivated channels have properties that are similar to those of expressed TRPs, leading some investigators to examine potential roles for TRP channels in myogenic constriction. For example, Welsh et al.33 demonstrated a role for TRPC6 in pressureinduced smooth muscle depolarization and vasoconstriction of rat cerebral resistance arteries. In this study, cerebral myocytes were patch clamped in the whole-cell conﬁguration. Cation currents were activated when cells were exposed to a hypotonic solution to induce membrane stretch. Downregulation of TRPC6 expression using antisense oligodeoxynucleotides attenuated swelling-activated currents in cerebral myocytes, suggesting that this channel is activated by mechanical stimulus in these cells. In isolated cerebral arteries, smooth muscle membrane depolarization and vasoconstriction induced by elevation of intraluminal pressure was also attenuated by TRPC6 antisense. These ﬁndings support the conclusion that membrane stretch activates a TRPC6-dependent depolarizing cation current that contributes to myogenic constriction of cerebral arteries. Although direct mechanosensitivity of the channel was not demonstrated, the authors speculate that membrane stretch activates PLC, thereby producing the second messenger DAG, which directly activates TRPC6.41 In support of this idea, Park et al.42 reported that mechanosensitive channels in vascular smooth muscle with properties similar to those of TRPC channels were inhibited by PLC blockade and activated by a DAG analogue. Furthermore, Slish et al.43 found that pressure-induced depolarization of cerebral artery myocytes was blocked by PLC inhibition. Two recent reports suggest that TRPM4, a member of the melastatin TRP family, may also contribute to stretch-induced depolarization of cerebral artery myocytes. In a preliminary study, Morita et al.44 reported activation of cation channels in cultured cells expressing cloned TRPM4 by applying negative pressure to the patch pipette. Similar stretch-activated channels were present in freshly isolated cerebral artery myocytes, suggesting that TRPM4 may be a mechanosensitive channel in these cells. Consistent with this proposal, Earley et al.45 reported that antisense-mediated downregulation of TRPM4 impaired pressure-induced depolarization and myogenic constriction of isolated cerebral arteries. Thus, it appears that both TRPC6 and TRPM4 are key components of the myogenic response in cerebral vascular myocytes and may participate in the autoregulation of cerebral blood ﬂow (see Figure 26.2). A study by Muraki et al.46 examined the possibility that TRPV2 is a mechanically activated ion channel in smooth muscle cells isolated from mouse aortas. The authors showed that hypotonic swelling of aortic myocytes activated a Ca2+ current that was not affected by inhibitors of L-type voltage-dependent calcium channels or pretreatment with caffeine to deplete intracellular Ca2+ stores. This swelling-activated current was blocked by ruthenium red, a nonselective inhibitor of TRPV channels, and by antisense-mediated suppression of TRPV2 expression. In addition, in cultured cells

368

TRP Ion Channel Function in Sensory Transduction

FIGURE 26.2 TRPC6, TRPM4, and TRPV2 channels are likely candidates as stretchactivated cation channels in vascular smooth muscle. These channels may be directly activated by membrane stretch or may be activated secondary to stretch activation of cell-signaling pathways (phospholipase C, cytoskeletal elements, integrin signaling) or other ion channels.

expressing cloned TRPV2, currents were activated by mechanical stretch (induced by applying negative pressure to the patch pipette) as well as by hypotonic stimuli. These ﬁndings suggest a potential role for TRPV2 in pressure-induced vasoconstrictor responses, although the effects of TRPV2 antisense on myogenic constriction or changes in smooth muscle membrane potential resulting from increased intravascular pressure in resistance arteries were not reported. Another recent report indicates that TRPC1 is a molecular candidate for the long-known mechanosensitive cation channel in oocytes.47 TRPC1 is clearly present in vascular smooth muscle cells,48 but no evidence has yet been presented to indicate a mechanosensitive role for TRPC1 in the vasculature. These reports suggest that multiple TRP channels expressed by vascular smooth muscle cells participate in pressure-induced depolarization and myogenic constriction of small arteries. Signaling pathways for channel activation during intraluminal pressure changes remain to be elucidated. However, it is clear that TRP channels are critical components of the myogenic response and likely play an important role in the autoregulation of blood ﬂow (see Figure 26.2).

STORE-OPERATED TRP CHANNEL ACTIVATION SMOOTH MUSCLE

IN

VASCULAR

In many cell types, depletion of calcium stores can trigger inﬂux of calcium via what have been termed store-operated channels (SOCs). This type of calcium inﬂux is thought to be important for maintaining normal function of the sarcoplasmic reticulum under a variety of conditions. The molecular identity of SOCs has been somewhat elusive, but evidence to suggest a role for TRP channels in this process is rather compelling. Recent studies suggest that TRPC1 is an integral component of store-operated Ca2+ channels in vascular smooth muscle cells. In isolated rabbit arteriolar smooth muscle cells, Xu and Beech48 depleted intracellular Ca2+ stores with thapsigargin in the presence of a voltage-gated Ca2+ channel blocker and “zero” extracellular Ca2+. When these cells were reintroduced to 1.5 mM extracellular Ca2+, an intracellular Ca2+ signal was recorded that was signiﬁcantly reduced in cells exposed to an

Functional Signiﬁcance of Transient Receptor Potential Channels

369

antibody that binds to an epitope near the proposed pore-forming region of TRPC1. Similarly, Sweeney et al.49 reported that both calcium entry and whole-cell cation currents induced by store depletion in proliferating human pulmonary artery smooth muscle cells are signiﬁcantly reduced following antisense-mediated TRPC1 downregulation. While blockade of TRPC1 reduces store-operated Ca2+ entry, overexpression of TRPC1 reportedly has the opposite effect and increases store-operated Ca2+ entry.50 In this study, rat pulmonary vascular rings were transfected with the human TRPC1 using an adenoviral vector. After 24 to 48 hours of organ culture, the rings were subjected to traditional protocols that elicit store-operated Ca2+ entry. Rings transfected with TRPC1 showed signiﬁcantly larger intracellular Ca2+ signals and greater active tension development when Ca2+ was reintroduced to the bath solution. Likewise, upregulation of TRPC1 expression during organ culture of rat cerebral arteries is associated with a substantial increase in store-operated Ca2+ entry,34 suggesting that TRPC1 forms store-operated Ca2+ inﬂux channels in vascular myocytes. Other studies support a role for either TRPC451 or TRPC652 in capacitative calcium entry and an associated proliferative response of human pulmonary artery smooth muscle cells. It appears that multiple TRPC channels may play important roles in the mechanism of calcium entry triggered in response to depletion of Ca2+ stores in vascular smooth muscle, and these Ca2+ responses may play a role in some vascular pathologies.

TRP CHANNEL FUNCTION IN VASCULAR ENDOTHELIAL CELLS PRESENCE

OF

TRP CHANNELS

IN THE

ENDOTHELIUM

There is good evidence derived from both molecular and histological approaches from multiple studies showing that TRP channels are widely expressed in endothelial cells. Although the TRP channel expression pattern in the endothelium appears to vary by species, all of the TRPC channel isoforms have been detected in endothelial cells.2 TRP channel isoforms from the two other major TRP subfamilies appear to be less abundant in endothelial cells. However, work by Fantozzi et al.53 and Wissenbach et al.54 illustrates that TRPM4 and TRPV4 are clearly present in endothelial cells.

TRP CHANNELS

AND

ENDOTHELIAL CELL CALCIUM ENTRY

Most TRP channels have substantial permeability to calcium ions. This may represent a signiﬁcant pathway for endothelial cell calcium inﬂux in response to various forms of stimulation, including receptor activation and shear stress, or following depletion of calcium stores. Evidence suggests that TRPC1 channels are involved in store-operated calcium entry in human pulmonary artery endothelial cells. A study by Brough et al.55 found that SOC entry induced by thapsigargin was decreased by about 40 percent in cultured endothelial cells treated with antisense oligonucleotides directed against TRPC1. Others have presented evidence showing that receptor activation triggers endothelial cell calcium entry, mediated at least in

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part by TRPC1. For instance, Antoniotti et al.56 observed that basic ﬁbroblast growth factor (bFGF) induced a sustained increase in cytoplasmic calcium levels in cultured bovine aortic endothelial cells. Addition of a polyclonal antibody against TRPC1 caused a 40 percent reversal of the bFGF-induced calcium increase, suggesting that the calcium inﬂux occurred through TRPC1 channels. In human mesenteric artery endothelial cells that prominently express TRPC1 channels, bradykinin activates a gadolinium-sensitive cation channel57 that represents a potential route for calcium entry. Although the molecular identity of the channel was not established in this study, its properties resemble those of expressed TRPC1 channels. Heterologously expressed TRPC3 channels are known to form both receptorand store-operated calcium channels.58 This also appears to be true for TRPC3 channels expressed by endothelial cells. Depletion of calcium stores in human umbilical vein endothelial cells induces a cation current that is abolished in cells expressing an N-terminal, dominant negative fragment of TRPC3.59 When human TRPC3 channels are heterologously expressed in bovine pulmonary endothelial cells, they are not activated by depletion of calcium stores but are activated by receptor ligands such as ATP and bradykinin.60 These ﬁndings clearly implicate TRPC3 as both an SOC entry pathway as well as a receptor-operated calcium entry channel and suggest that TRPC3 channels are centrally involved in endotheliumdependent regulation of vascular function (see Figure 26.3). TRPC4 knockout mice have provided a powerful tool for demonstrating functional roles for this channel. Studies using these mice suggest that TRPC4 channels expressed by endothelial cells appear to be important in store-operated and agonistinduced calcium entry and endothelium-dependent vasodilation. Store-operated calcium currents in aortic endothelial cells are abolished in TRPC4 knockout mice.61 Further, agonist-induced endothelial calcium entry and endothelium-dependent vasodilation are also greatly suppressed in these mice. Similarly, calcium entry induced by thrombin or by activation of the PAR-1 receptor in pulmonary endothelial

Receptor-activated TRPC3, TRPC4, or TRPV4 channels Store-operated TRPC1 or TRPC3 channels Ca2+

Vasodilation

FIGURE 26.3 Endothelial TRP channels may contribute to the regulation of calcium entry and activity of endothelial vasodilator factors.

Functional Signiﬁcance of Transient Receptor Potential Channels

371

cells is absent in TRPC4 knockout mice.62 It has been suggested that activation of store-operated, endothelial cell calcium entry via TRPC4 channels is regulated through physical interactions of TRPC4 channels with the actin cytoskeleton, perhaps as part of a larger protein complex that could include TRPC1 as well.63 Endothelial cell cation channels having properties very similar to those of cloned TRPV4 channels are activated by 5,6 EET,64,65 a putative endothelium-derived hyperpolarizing factor.66 Channel activation by 5,6 EET is accompanied by a substantial increase in endothelial cell calcium, implying that endothelial TRPV4 channels may contribute to the vasodilator effects of endogenous arachidonic acid metabolites. Thus TRPV4 channels located both in endothelial cells and in smooth muscle (see above) appear to contribute to the vasodilator response to EETs.

TRP CHANNELS IN THE PROLIFERATIVE RESPONSE ENDOTHELIUM TO HYPOXIA

OF THE

Endothelial cells proliferate in response to hypoxic exposure. Fantozzi et al.53 have presented evidence suggesting a role for endothelial TRP channels in this proliferative response in human pulmonary artery endothelial cells. Proliferation is associated with an increase in the expression of AP-1 receptors, elevated store-operated calcium entry, and enhanced TRPC4 channel expression. The increase in AP-1 receptor number following hypoxic exposure is greatly reduced in endothelial cells treated with TRPC antisense oligonucleotides, suggesting a linkage between TRPC4 channels and the proliferative response of pulmonary endothelial cells to hypoxia.

TRP CHANNELS

AND

REGULATION

OF

CAPILLARY PERMEABILITY

A key role for TRPC4 channels in the regulation of endothelial capillary permeability has been recently proposed.67 Activation of thrombin receptors or the proteinaseactivated receptor-1 (PAR-1) in wild-type pulmonary endothelial cells induces a lanthanum-sensitive calcium inﬂux and an associated increase in endothelial capillary permeability.62 These responses were reduced by about 50 percent in endothelial cells from TRPC4 knockout versus wild-type mice, suggesting that TRPC4 channels are critical mediators of thrombin and PAR-1-induced Ca2+ inﬂux.

ROLE OF TRP CHANNELS IN ENDOTHELIAL CELL RESPONSE TEMPERATURE CHANGE

TO

TRPV channels are involved in cellular responses to environmental stimuli such as changes in osmolarity and temperature.4 TRPV4 channels are highly expressed in endothelial cells, and a recent study suggests that these channels may be involved in regulation of vascular function in response to changes in ambient temperature.68 These ﬁndings demonstrate that the properties of native thermosensitive cation channels in aortic endothelial cells were indistinguishable from those of heterologously expressed TRPV4 channels. The authors propose that altered calcium inﬂux through TRPV4 at different temperatures could modulate nitric oxide production to induce vasoconstriction at lower temperatures and vasodilation at higher temperatures.

372

TRP Ion Channel Function in Sensory Transduction

TRP CHANNELS

AS

SENSORS

OF

ENDOTHELIAL OXIDATIVE STRESS

TRPC3 channels may be involved in the endothelial cell response to oxidative stress.69 One study reports that oxidative stress of cultured porcine aortic endothelial cells activates a nonselective cation current and depolarizes the endothelial cells. The oxidant-induced current is abolished in cells expressing an N-terminal, dominant negative TRPC3 fragment, clearly implicating TRPC3 in the response.

SUMMARY AND FUTURE PERSPECTIVES Growing evidence demonstrates that TRP channel proteins form functional nonselective cation channels in vascular smooth muscle and endothelial cells (Table 26.1). These channels appear to be activated by various stimuli including mechanical perturbations, receptor activation, calcium store depletion, or changes in temperature, and the channels may be involved in multiple cellular processes in the vasculature. The speciﬁc roles of these channels in normal and pathologically impaired vascular function are only just beginning to come to light but include neurotransmission, hormonal control of the vasculature, nitric oxide production, myogenic tone and autoregulation of blood ﬂow, thermoregulation, responses to oxidative stress, and cellular proliferative activity. Further research toward understanding the interactions of TRP channels with other proteins and signaling mechanisms is clearly needed in order to understand the functional signiﬁcance of TRP channels in vascular smooth muscle and endothelial cells.

REFERENCES 1. Albert, A.P. and Large, W.A., Signal transduction pathways and gating mechanisms of native TRP-like cation channels in vascular myocytes, J. Physiol. 570, 45–51, 2005. 2. Yao, X. and Garland, C.J., Recent developments in vascular endothelial cell transient receptor potential channels, Circ. Res. 97 (9), 853–63, 2005. 3. Beech, D.J., Muraki, K., and Flemming, R., Non-selective cationic channels of smooth muscle and the mammalian homologues of Drosophila TRP, J. Physiol. 559 (Pt. 3), 685–706, 2004. 4. Clapham, D.E., TRP channels as cellular sensors, Nature 426 (6966), 517–24, 2003. 5. Xu, X.Z., Li, H.S., Guggino, W.B., and Montell, C., Coassembly of TRP and TRPL produces a distinct store-operated conductance, Cell 89 (7), 1155–64, 1997. 6. Hofmann, T., Schaefer, M., Schultz, G., and Gudermann, T., Subunit composition of mammalian transient receptor potential channels in living cells, Proc. Natl. Acad. Sci. USA 99 (11), 7461–66, 2002. 7. Strubing, C., Krapivinsky, G., Krapivinsky, L., and Clapham, D.E., Formation of novel TRPC channels by complex subunit interactions in embryonic brain, J. Biol. Chem. 278 (40), 39014–19, 2003. 8. Strubing, C., Krapivinsky, G., Krapivinsky, L., and Clapham, D.E., TRPC1 and TRPC5 form a novel cation channel in mammalian brain, Neuron 29 (3), 645–55, 2001. 9. Walker, R.L., Hume, J.R., and Horowitz, B., Differential expression and alternative splicing of TRP channel genes in smooth muscles, Am. J. Physiol. Cell Physiol. 280 (5), C1184–92, 2001.

Functional Signiﬁcance of Transient Receptor Potential Channels

373

10. Lee, M.W. and Severson, D.L., Signal transduction in vascular smooth muscle: diacylglycerol second messengers and PKC action, Am. J. Physiol. 267 (3 Pt. 1), C659–78, 1994. 11. Berridge, M.J., Lipp, P., and Bootman, M.D., The versatility and universality of calcium signalling, Nat. Rev. Mol. Cell Biol. 1 (1), 11–21, 2000. 12. Putney, J.W., Jr., A model for receptor-regulated calcium entry, Cell Calcium 7 (1), 1–12, 1986. 13. Chen, X.L. and Rembold, C.M., Phenylephrine contracts rat tail artery by one electromechanical and three pharmacomechanical mechanisms, Am. J. Physiol. 268 (1 Pt. 2), H74–81, 1995. 14. Nelson, M.T., Patlak, J.B., Worley, J.F., and Standen, N.B., Calcium channels, potassium channels, and voltage dependence of arterial smooth muscle tone, Am. J. Physiol. 259 (1 Pt. 1), C3–18, 1990. 15. Karaki, H., Ozaki, H., Hori, M., Mitsui-Saito, M., Amano, K., Harada, K., Miyamoto, S., Nakazawa, H., Won, K.J., and Sato, K., Calcium movements, distribution, and functions in smooth muscle, Pharmacol. Rev. 49 (2), 157–230, 1997. 16. Byrne, N.G. and Large, W.A., Membrane ionic mechanisms activated by noradrenaline in cells isolated from the rabbit portal vein, J. Physiol. 404, 557–73, 1988. 17. Chen, C. and Wagoner, P.K., Endothelin induces a nonselective cation current in vascular smooth muscle cells, Circ. Res. 69 (2), 447–54, 1991. 18. Helliwell, R.M. and Large, W.A., Alpha 1-adrenoceptor activation of a non-selective cation current in rabbit portal vein by 1,2-diacyl-sn-glycerol, J. Physiol. 499 (Pt. 2), 417–28, 1997. 19. Nakajima, T., Hazama, H., Hamada, E., Wu, S.N., Igarashi, K., Yamashita, T., Seyama, Y., Omata, M., and Kurachi, Y., Endothelin-1 and vasopressin activate Ca(2+)permeable non-selective cation channels in aortic smooth muscle cells: mechanism of receptor-mediated Ca2+ inﬂux, J. Mol. Cell Cardiol. 28 (4), 707–22, 1996. 20. Montell, C., TRP trapped in ﬂy signaling web, Curr. Opin. Neurobiol. 8 (3), 389–97, 1998. 21. Hardie, R.C. and Minke, B., Phosphoinositide-mediated phototransduction in Drosophila photoreceptors: the role of Ca2+ and trp, Cell Calcium 18 (4), 256–74, 1995. 22. Yu, Y., Fantozzi, I., Remillard, C.V., Landsberg, J.W., Kunichika, N., Platoshyn, O., Tigno, D.D., Thistlethwaite, P.A., Rubin, L.J., and Yuan, J.X., Enhanced expression of transient receptor potential channels in idiopathic pulmonary arterial hypertension, Proc. Natl. Acad. Sci. USA 101 (38), 13861–66, 2004. 23. McDaniel, S.S., Platoshyn, O., Wang, J., Yu, Y., Sweeney, M., Krick, S., Rubin, L.J., and Yuan, J.X., Capacitative Ca(2+) entry in agonist-induced pulmonary vasoconstriction, Am. J. Physiol. Lung Cell Mol. Physiol. 280 (5), L870–L880, 2001. 24. Clapham, D.E., Runnels, L.W., and Strubing, C., The TRP ion channel family, Nat. Rev. Neurosci. 2 (6), 387–96, 2001. 25. Harteneck, C., Plant, T.D., and Schultz, G., From worm to man: three subfamilies of TRP channels, Trends Neurosci. 23 (4), 159–66, 2000. 26. Lintschinger, B., Balzer-Geldsetzer, M., Baskaran, T., Graier, W.F., Romanin, C., Zhu, M.X., and Groschner, K., Coassembly of Trp1 and Trp3 proteins generates diacylglycerol- and Ca2+-sensitive cation channels, J. Biol. Chem. 275 (36), 27799–805, 2000. 27. Wu, X., Babnigg, G., and Villereal, M.L., Functional signiﬁcance of human trp1 and trp3 in store-operated Ca(2+) entry in HEK-293 cells, Am. J. Physiol. Cell Physiol. 278 (3), C526–36, 2000.

374

TRP Ion Channel Function in Sensory Transduction

28. Inoue, R., Okada, T., Onoue, H., Hara, Y., Shimizu, S., Naitoh, S., Ito, Y., and Mori, Y., The transient receptor potential protein homologue TRP6 is the essential component of vascular alpha(1)-adrenoceptor-activated Ca(2+)-permeable cation channel, Circ. Res. 88 (3), 325–32, 2001. 29. Jung, S., Strotmann, R., Schultz, G., and Plant, T.D., TRPC6 is a candidate channel involved in receptor-stimulated cation currents in A7r5 smooth muscle cells, Am. J. Physiol. Cell Physiol. 282 (2), C347–59, 2002. 30. Welsh, D.G. and Brayden, J.E., Mechanisms of coronary artery depolarization by uridine triphosphate, Am. J. Physiol. Heart Circ. Physiol. 280 (6), H2545–53, 2001. 31. Reading, S.A., Earley, S., Waldron, B.J., Welsh, D.G., and Brayden, J.E., TRPC3 mediates pyrimidine receptor-induced depolarization of cerebral arteries, Am. J. Physiol. Heart Circ. Physiol. 288 (5), H2055–61, 2005. 32. Reading, S.A., Lutz, K.E., and Brayden, J.E., Ca2+ inﬂux through TRPC3 channels contributes to agonist-induced constriction of cerebral arteries., FASEB J. 19 (5), A1625, 2005. 33. Welsh, D.G., Morielli, A.D., Nelson, M.T., and Brayden, J.E., Transient receptor potential channels regulate myogenic tone of resistance arteries, Circ. Res. 90 (3), 248–50, 2002. 34. Bergdahl, A., Gomez, M.F., Wihlborg, A.K., Erlinge, D., Eyjolfson, A., Xu, S.Z., Beech, D.J., Dreja, K., and Hellstrand, P., Plasticity of TRPC expression in arterial smooth muscle: correlation with store-operated Ca2+ entry, Am. J. Physiol. Cell Physiol. 288 (4), C872–80, 2005. 35. Lin, M.J., Leung, G.P., Zhang, W.M., Yang, X.R., Yip, K.P., Tse, C.M., and Sham, J.S., Chronic hypoxia-induced upregulation of store-operated and receptor-operated Ca2+ channels in pulmonary arterial smooth muscle cells: a novel mechanism of hypoxic pulmonary hypertension, Circ. Res. 95 (5), 496–505, 2004. 36. Earley, S., Heppner, T.J., Nelson, M.T., and Brayden, J.E., TRPV4 forms a novel Ca2+ signaling complex with ryanodine receptors and BKCa channels, Circ. Res. 97 (12), 1270–79, 2005. 37. Bayliss, W.M., On the local reactions of the arterial wall to changes of internal pressure, J. Physiol. (London) 28, 220–31, 1902. 38. Davis, M.J. and Hill, M.A., Signaling mechanisms underlying the vascular myogenic response, Physiol. Rev. 79 (2), 387–423, 1999. 39. Knot, H.J. and Nelson, M.T., Regulation of arterial diameter and wall [Ca2+] in cerebral arteries of rat by membrane potential and intravascular pressure, J. Physiol. 508 (Pt. 1), 199–209, 1998. 40. Harder, D.R., Pressure-dependent membrane depolarization in cat middle cerebral artery, Circ. Res. 55 (2), 197–202, 1984. 41. Hofmann, T., Obukhov, A.G., Schaefer, M., Harteneck, C., Gudermann, T., and Schultz, G., Direct activation of human TRPC6 and TRPC3 channels by diacylglycerol, Nature 397 (6716), 259–63, 1999. 42. Park, K.S., Kim, Y., Lee, Y.H., Earm, Y.E., and Ho, W.K., Mechanosensitive cation channels in arterial smooth muscle cells are activated by diacylglycerol and inhibited by phospholipase C inhibitor, Circ. Res. 93 (6), 557–64, 2003. 43. Slish, D.F., Welsh, D.G., and Brayden, J.E., Diacylglycerol and protein kinase C activate cation channels involved in myogenic tone, Am. J. Physiol. Heart Circ. Physiol. 283 (6), H2196–201, 2002. 44. Morita, H., Honda, A., Nelson, M.T., and Brayden, J.E., Stretch activates Ca2+sensitive, non-selective cation channels in smooth muscle cells from cerebral arteries, FASEB J. 17 (4), A64 (Abstract), 2003.

Functional Signiﬁcance of Transient Receptor Potential Channels

375

45. Earley, S., Waldron, B.J., and Brayden, J.E., Critical role for transient receptor potential channel TRPM4 in myogenic constriction of cerebral arteries, Circ. Res. 95 (9), 922–29, 2004. 46. Muraki, K., Iwata, Y., Katanosaka, Y., Ito, T., Ohya, S., Shigekawa, M., and Imaizumi, Y., TRPV2 is a component of osmotically sensitive cation channels in murine aortic myocytes, Circ. Res. 93 (9), 829–38, 2003. 47. Maroto, R., Raso, A., Wood, T.G., Kurosky, A., Martinac, B., and Hamill, O.P., TRPC1 forms the stretch-activated cation channel in vertebrate cells, Nat. Cell Biol. 7 (2), 179–85, 2005. 48. Xu, S.Z. and Beech, D.J., TrpC1 is a membrane-spanning subunit of store-operated Ca(2+) channels in native vascular smooth muscle cells, Circ. Res. 88 (1), 84–87, 2001. 49. Sweeney, M., Yu, Y., Platoshyn, O., Zhang, S., McDaniel, S.S., and Yuan, J.X., Inhibition of endogenous TRP1 decreases capacitative Ca2+ entry and attenuates pulmonary artery smooth muscle cell proliferation, Am. J. Physiol. Lung Cell Mol. Physiol. 283 (1), L144–55, 2002. 50. Kunichika, N., Yu, Y., Remillard, C.V., Platoshyn, O., Zhang, S., and Yuan, J.X., Overexpression of TRPC1 enhances pulmonary vasoconstriction induced by capacitative Ca2+ entry, Am. J. Physiol. Lung Cell Mol. Physiol. 287 (5), L962–99, 2004. 51. Zhang, S., Remillard, C.V., Fantozzi, I., and Yuan, J.X., ATP-induced mitogenesis is mediated by cyclic AMP response element-binding protein-enhanced TRPC4 expression and activity in human pulmonary artery smooth muscle cells, Am. J. Physiol. Cell Physiol. 287 (5), C1192–201, 2004. 52. Yu, Y., Sweeney, M., Zhang, S., Platoshyn, O., Landsberg, J., Rothman, A., and Yuan, J.X., PDGF stimulates pulmonary vascular smooth muscle cell proliferation by upregulating TRPC6 expression, Am. J. Physiol. Cell Physiol. 284 (2), C316–30, 2003. 53. Fantozzi, I., Zhang, S., Platoshyn, O., Remillard, C.V., Cowling, R.T., and Yuan, J.X., Hypoxia increases AP-1 binding activity by enhancing capacitative Ca2+ entry in human pulmonary artery endothelial cells, Am. J. Physiol. Lung Cell Mol. Physiol. 285 (6), L1233–45, 2003. 54. Wissenbach, U., Bodding, M., Freichel, M., and Flockerzi, V., Trp12, a novel Trprelated protein from kidney, FEBS Lett. 485 (2–3), 127–34, 2000. 55. Brough, G.H., Wu, S., Ciofﬁ, D., Moore, T.M., Li, M., Dean, N., and Stevens, T., Contribution of endogenously expressed Trp1 to a Ca2+-selective, store-operated Ca2+ entry pathway, FASEB J. 15 (10), 1727–38, 2001. 56. Antoniotti, S., Lovisolo, D., Fiorio Pla, A., and Munaron, L., Expression and functional role of bTRPC1 channels in native endothelial cells, FEBS Lett. 510 (3), 189–95, 2002. 57. Kohler, R., Brakemeier, S., Kuhn, M., Degenhardt, C., Buhr, H., Pries, A., and Hoyer, J., Expression of ryanodine receptor type 3 and TRP channels in endothelial cells: comparison of in situ and cultured human endothelial cells, Cardiovasc. Res. 51 (1), 160–68, 2001. 58. Venkatachalam, K., van Rossum, D.B., Patterson, R.L., Ma, H.T., and Gill, D.L., The cellular and molecular basis of store-operated calcium entry, Nat. Cell Biol. 4 (11), E263–72, 2002. 59. Groschner, K., Hingel, S., Lintschinger, B., Balzer, M., Romanin, C., Zhu, X., and Schreibmayer, W., Trp proteins form store-operated cation channels in human vascular endothelial cells, FEBS Lett. 437 (1–2), 101–6, 1998. 60. Kamouchi, M., Philipp, S., Flockerzi, V., Wissenbach, U., Mamin, A., Raeymaekers, L., Eggermont, J., Droogmans, G., and Nilius, B., Properties of heterologously expressed

376

61.

62.

63. 64.

65.

66.

67.

68.

69.

70.

71.

TRP Ion Channel Function in Sensory Transduction hTRP3 channels in bovine pulmonary artery endothelial cells, J. Physiol. 518 (Pt. 2), 345–58, 1999. Freichel, M., Suh, S. H., Pfeifer, A., Schweig, U., Trost, C., Weissgerber, P., Biel, M., Philipp, S., Freise, D., Droogmans, G., Hofmann, F., Flockerzi, V., and Nilius, B., Lack of an endothelial store-operated Ca2+ current impairs agonist-dependent vasorelaxation in TRP4-/- mice, Nat. Cell Biol. 3 (2), 121–27, 2001. Tiruppathi, C., Freichel, M., Vogel, S.M., Paria, B.C., Mehta, D., Flockerzi, V., and Malik, A.B., Impairment of store-operated Ca2+ entry in TRPC4(-/-) mice interferes with increase in lung microvascular permeability, Circ. Res. 91 (1), 70–76, 2002. Ciofﬁ, D.L., Wu, S., and Stevens, T., On the endothelial cell I(SOC), Cell Calcium 33 (5–6), 323–36, 2003. Watanabe, H., Vriens, J., Prenen, J., Droogmans, G., Voets, T., and Nilius, B., Anandamide and arachidonic acid use epoxyeicosatrienoic acids to activate TRPV4 channels, Nature 424 (6947), 434–38, 2003. Vriens, J., Owsianik, G., Fisslthaler, B., Suzuki, M., Janssens, A., Voets, T., Morisseau, C., Hammock, B.D., Fleming, I., Busse, R., and Nilius, B., Modulation of the Ca2+ permeable cation channel TRPV4 by cytochrome P450 epoxygenases in vascular endothelium, Circ. Res. 97 (9), 908–15, 2005. Campbell, W.B. and Harder, D.R., Endothelium-derived hyperpolarizing factors and vascular cytochrome P450 metabolites of arachidonic acid in the regulation of tone, Circ. Res. 84 (4), 484–88, 1999. Tiruppathi, C., Minshall, R.D., Paria, B.C., Vogel, S.M., and Malik, A.B., Role of Ca2+ signaling in the regulation of endothelial permeability, Vascul. Pharmacol. 39 (4–5), 173–85, 2002. Watanabe, H., Vriens, J., Suh, S.H., Benham, C.D., Droogmans, G., and Nilius, B., Heat-evoked activation of TRPV4 channels in a HEK293 cell expression system and in native mouse aorta endothelial cells, J. Biol. Chem. 277 (49), 47044–51, 2002. Balzer, M., Lintschinger, B., and Groschner, K., Evidence for a role of Trp proteins in the oxidative stress-induced membrane conductances of porcine aortic endothelial cells, Cardiovasc. Res. 42 (2), 543–49, 1999. Yip, H., Chan, W.Y., Leung, P.C., Kwan, H.Y., Liu, C., Huang, Y., Michel, V., Yew, D.T., and Yao, X., Expression of TRPC homologs in endothelial cells and smooth muscle layers of human arteries, Histochem. Cell Biol. 122 (6), 553–61, 2004. Facemire, C.S., Mohler, P.J., and Arendshorst, W.J., Expression and relative abundance of short transient receptor potential channels in the rat renal microcirculation, Am. J. Physiol. Renal Physiol. 286 (3), F546–51, 2004.

27

Role of TRPV4 in the Mechanotransduction of Shear Stress in Endothelial Cells Ralf Köhler and Joachim Hoyer Philipps University

CONTENTS Ca2+ and Endothelial Function ..............................................................................378 TRPs in the Endothelium ......................................................................................379 TRPV4 in the Endothelium...................................................................................380 TRPV4 and Endothelium-Dependent Vasodilatation............................................382 TRPV4-Mediated Vasodilatation is Nitric Oxide Dependent in Carotid Arteries .................................................................................................383 A Functional Role of TRPV4 in Endothelial Mechanotransduction?..................383 Endothelial TRP Channels and Cardiovascular Disease ......................................385 Do Endothelial Ca2+-Permeable Cation Channels Like TRPV4 Represent Novel Pharmacotherapeutic Targets for Anti-Hypertensive Therapy? ................................................................................................................386 References..............................................................................................................386 The endothelium is a highly specialized multifunctional cell monolayer between blood and tissue. It regulates a variety of vascular functions such as the passage of macromolecules and oxygen supply to organs and tissues, of immune responses, of angiogenesis, and of vascular remodeling. An additional and overall important role of the arterial endothelium is the control of the contractile state of the vascular smooth muscle and thus systemic blood pressure by the release of vasoactive factors. Moreover, endothelial dysfunction contributes to several cardiovascular pathologies such as arteriosclerosis, restenosis disease, vasculitis, or hypertension. The crucial role of the endothelium in the control of vascular tone was ﬁrst demonstrated by Furchgott and Zawadzki in 1980 [1]. In this pioneer work, they demonstrated that following stimulation with acetylcholine, the endothelium releases a short-lived factor that relaxes the vascular smooth muscle. This factor was later

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identiﬁed as nitric oxide. In addition to humoral stimulation, the endothelium also controls vasodilatation in response to increased hemodynamic forces (i.e., increased shear stress exerted by streaming blood) [2]. This is an important mechanism by which the endothelium controls adequate organ perfusion and protects vessel walls against mechanical damage.

CA2+ AND ENDOTHELIAL FUNCTION In response to classical agonists such as acetylcholine and bradykinin as well as to hemodynamic stimuli, the endothelium produces in principle three types of vasodilating factors: nitric oxide (NO), prostacyclin (PGI2), and the endothelium-derived hyperpolarizing factor (EDHF) (Figure 27.1). In contrast to nitric oxide and PGI2, the EDHF is not considered a chemical vasodilator [3,4] but rather an electrical phenomenon in which an initial endothelial hyperpolarization spreads to the vascular smooth muscle via a myoendothelial gap junction, closing L-type voltage-gated Ca2+ channels and thus vasorelaxation. The synthesis of all three vasodilators is Ca2+ dependent. The endothelial nitric oxide synthase is activated in a Ca2+/calmodulin-dependent fashion [5], and also PGI2 synthesis by cyclooxgenase-1 requires the Ca2+-dependent release of arachidonic acid by phospholipase-A2. The generation of the EDHF signaling is Ca2+ dependent

FIGURE 27.1 Schematic illustration of the hypothetical role of TRPC and mechanosensitive TRPV4 channels in endothelial function. 4αPDD, 4α-phorbol-12,13-didecanoate; AA, arachidonic acid; EETs, epoxyeicosatrienoic acids; COX, cyclooxygenase-1; CYP, cytochrome P450 epoxygenase; DAG, diacylglycerol; EDHF, endothelium-derived hyperpolarizing factor; ER, endoplasmic reticulum; KCa3.1 and KCa2.3, endothelial Ca2+-activated K+-channels; IP3, inositoltrisphosphate; MSC, mechanosensitive cation channels; NOS, endothelial NOsynthase; NO, nitric oxide; PGI2, prostacyclin; RuR, ruthenium red.

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because the initial endothelial hyperpolarization is caused by opening endothelial Ca2+-activated K+ channels [4,6–10]. Upon endothelial stimulation by binding of classical agonists to their G-proteincoupled receptors, the intracellular Ca2+ concentration ([Ca2+]i) increases rapidly within seconds due to an IP3-mediated Ca2+ release from internal stores. Following the initial Ca2+ peak, [Ca2+]i remains elevated for up to several minutes, due to Ca2+ inﬂux from the extracellular space. Ca2+-permeable cation channels located in the plasma membrane play an important role in this plateau phase of elevated [Ca2+]i as they provide the Ca2+-inﬂux pathway [11]. Besides the well-documented Ca2+ mobilization following receptor stimulation, stimulation of the endothelium by hemodynamic stimuli (i.e., shear stress) also results in an increase in [Ca2+]i. Such a shear stress–induced Ca2+ mobilization has been observed following stimulation in endothelial cells in vitro [9,12] as well as in the endothelium in intact vessel preparations [13]. This ﬂow/shear stress–induced increase in [Ca2+]i is thought to involve Ca2+ inﬂux through mechanosensitive Ca2+-permeable cation channels (MSC), which are believed to act as mechanosensors in recognizing alterations of hemodynamic forces, as well as the Ca2+ release from internal IP3- [14] and ryanodine-sensitive stores [9]. The Ca2+ inﬂux through MSC- and Ca2+-release events result in [Ca2+]i oscillations with a complex spatial and temporal pattern [9,12]. Frequency and amplitude modulation of such [Ca2+]i ﬂuctuations is thought to ﬁne-tune the synthesis of endothelial vasodilators depending on the degree of mechanical stimulation [9]. At the cell membrane level, MSC has been identiﬁed in EC in vitro [11,15] and in situ [16,17] as mostly stretch-activated channels (SAC) based on their activation by applying negative pressure to the patch pipette in single-channel patch-clamp recordings. However, molecular identity (i.e., the MSC/SAC encoding gene[s] as well as the intrinsic molecular determinants of mechanosensitivity) is still elusive. However, the recent identiﬁcation of cation channels of the TRP gene family with mechanosensitive properties may bring some progress in this ﬁeld [18,19].

TRPS IN THE ENDOTHELIUM In recent years, the mammalian genes encoding for Ca2+-permeable cation channels were identiﬁed as mammalian homologues of the “transient receptor potential” trp gene expressed in photoreceptors of Drosophila [20]; meanwhile, expressions of several members of the TRP superfamily of cation channels have been shown in endothelial cells (EC) of humans and other species (for review, see reference 21). Within the diverse subfamilies of TRP cation channels, EC express several members of the canonical TRP subfamily (TRPC): TRPC1, TRPC3, TRPC4, and TRPC6 with expression patterns varying within different species. Homomeric or heteromeric TRPC channel complexes are principally thought to serve as Ca2+ inﬂux channels after receptor activation or store depletion (Figure 27.1) [21,22]. Of the endothelial TRPCs, particularly TRPC4 has been shown to contribute to endothelial-dependent vasodilatation as concluded from an altered vasorelaxation response in TRPC4 knockout mice [23]. Concerning the other TRP subfamilies, the melastatin (TRPM) and vanilloid (TRPV) subfamilies (e.g., TRPM4 and TRPV4) have also been reported to be endothelial TRPs [21,24].

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The functional role of TRPM4 in the endothelium is unclear. In addition to its regulation by ATP- and PKC-mediated phosphorylation, this channel is itself activated by increases in intracellular Ca2+ in a Ca2+/calmodulin-dependent fashion, but it is not Ca2+ permeable [21]. The depolarizing Na+ current through this channel leads to membrane depolarization. Possibly, this channel is needed (in terms of a negative feedback mechanism) to counteract prolonged membrane hyperpolarization, which is caused by open Ca2+-activated K+ channels. TRPV4 was ﬁrst identiﬁed in the endothelium of mouse aorta by Bernd Nilius’s group [24]. Thus far, it appears that this channel is the only member of the TRPV subfamily expressed in the endothelium. TRPV4 was also identiﬁed by us in the endothelium of the carotid artery of the rat. The precise role of this TRPV subtype in endothelial function is not elucidated so far. However, in this chapter we provide new insight into how TRPV4 contributes to endothelial function and endotheliumdependent vasodilatation.

TRPV4 IN THE ENDOTHELIUM Within this diverse expression pattern of TRP channels in the endothelium, TRPV4 is especially interesting because of its moderately high Ca2+ permeability [24,25], which would offer a signiﬁcant Ca2+-inﬂux pathway (Figure 27.1). Unlike TRPC channels, TRPV4 is not considered a classical receptor/second-messenger–activated channel or store-operated channel [20], but it exhibits extremely diverse gating behavior upon physical and chemical stimuli and importantly mechanical and osmotic challenges [26]. Due to this mechano- and osmosensitivity, the new term mechano- and osmo- TRP was assigned to this TRPV4 [19]. Moreover, the physiological importance of this channel and of OSM-9, the TRPV homologue in invertebrate C. elegans, was highlighted by the ﬁnding of a disturbed mechanosensation and osmoregulation in TRPV4 knockout mice [27] and the impaired mechanosensation, olfaction, and osmoregulation in C. elegans with a mutant osm-9-gene [28]. The detailed biophysical properties of this channel and its molecular mechanism of activation were outlined in other chapters of this book. After a brief summary of general functional features of endothelial TRPV4, we focus in this chapter on the potential physiological roles of TRPV4 in the endothelium and in the mechanism of endothelial control of vascular tone. Similar to TRPV4 in heterologous expression systems, TRPV4 in aortic endothelial cells in mice [24,29] and in the EC of the rat carotid artery is activated by moderate warmth (>27°C), cell swelling in response and hypotonic stress (HTS), and pharmacologically by the non-PKC-activating phorbol ester, 4α-phorbol-12,13didecanoate (4αPDD), leading to a considerable increase in endothelial [Ca2+]i (Figure 27.2A–C). Single-cell-RT-PCR analysis of the TRPV mRNA expression pattern in the in situ EC of carotid arteries revealed TRPV4 expression, but none of the other closely related TRPV1–3. Within the vascular wall of the rat carotid artery, TRPV4 expression seems to be limited to the endothelium because mRNA expression was not detectable in the vascular smooth muscle cells in the carotid artery. However, it should be noted that TRPV4 expression and channel functions were recently detected in the smooth muscle of cerebral arteries of rats [30], in which

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FIGURE 27.2 Electrophysiological properties of TRPV4 currents in RCAEC of rat carotid artery in situ. (A) Representative whole-cell recording of activation of cation currents by 4α-phorbol-12,13-didecanoate (4αPDD, 1 μM). Nonpermeable NMDG as the only cation in the bath solution abolishes currents at negative membrane potentials. Voltage-dependent inhibition of TRPV4 currents by ruthenium red (RuR, 1 μM). (B) Time course of 4αPDDinduced currents at [Ca2+] out (1 mM). (C) Increase in [Ca2+]i by 150 nM above basal levels following activation of TRPV4 by 4αPDD in freshly isolated RAEC. (D) TRPV4 and endothelium-dependent vasodilatation. 4αPDD (1 μM) induced strong vasodilatation in carotid arteries with a functionally active endothelium. Vasodilatation to 4αPDD is suppressed by the TRPV4 blocker RuR (1 μM). Note that the vasodilatory response elicited by 4αPDD is of similar magnitude as that induced by ACh (1 μM). (E) EDHF-type vasodilatation induced by ACh (1 μM), endothelial KCa channel opener 1-EBIO (100 μM), but not by 4αPDD (1 μM) in the presence of the NO-synthase inhibitor L-NNA (100 μM) and the cyclooxygenase inhibitor INDO (10 μM). (F) Vasodilatation in response to increased shear stress/viscosity (5 percent dextran) and inhibitory effects of RuR (1 μM) and L-NNA and INDO.

this channel forms a functional complex with the large-conductance Ca2+activated K+ channel (BKCa), which modulates vascular smooth muscle membrane potential. This also indicates that TRPV4 expression may be heterogeneous within different vascular beds. With respect to mechanism of TRPV4 activation in the endothelium, arachidonic acid (AA) has been shown to mediate HTS-induced TRPV4 activation similar to ﬁndings in other cell types and in heterologous expression systems [31,32]. A more recent study in aortic ECs from mice suggests that arachidonic acid itself might not

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be the endogenous activator of the channel. But conversion of arachidonic acid to 5,6 epoxyeicosatrienoic acid (5,6 EET) and 8,9 epoxyeicosatrienoic acid (8,9 EET) mediated by cytochrome P450 epoxygenases leads to channel activation [33]. Thus, the authors proposed that EETs serve as the endogenous activators of TRPV4. This ﬁnding is highly relevant and offers some exciting insight into the putative role of TRPV4 in the endothelial function because EETs have been suggested to be molecular candidates for the EDHF (the third major vasodilating factor) in some vessels or to facilitate EDHF signaling by intra-endothelial mechanisms [4,33,34]. Therefore, EET activation of TRPV4 and subsequent Ca2+ entry may play a role in the generation or ampliﬁcation of EDHF signaling after receptor stimulation.

TRPV4 AND ENDOTHELIUM-DEPENDENT VASODILATATION The mechanosensitivity of TRPV4 may point to a role of the channel as an endothelial mechanosensor and thus in the mechanisms of ﬂow- or shear stress–induced vasodilatation. It is noteworthy that a shear stress–induced TRPV4-mediated Ca2+ entry has also been shown in heterologous expression systems and renal tubular epithelial cells [35,36]. As mentioned before, such ﬂow- and shear stress–induced Ca2+ signals have also been observed in the EC in vitro [9,12] and in the endothelium in intact vessel preparations [13]. The inhibition of MSC by the rather nonselective lanthanide gadolinium (Gd3+) abolishes this ﬂow- and shear stress–induced Ca2+ mobilization [9]. As already stated, the molecular identity of this shear stress–activated Ca2+-permeable MSC is still unknown. But in keeping with the mechanosensitivity of TRPV4, this channel might be a good molecular candidate for the shear stress–activated Ca2+-entry channel in the endothelium. In our own studies, we characterized the functional role of TRPV4 channels in the endothelium of rat carotid arteries by pressure myography, and we determined vasodilatory responses after pharmacological activation of the channel by 4αPDD and after increasing shear stress. Our myograph experiments revealed that pharmacological opening of TRPV4 by 4αPDD caused a robust vasodilatation in the carotid artery (Figure 27.2D). This 4αPDD-induced vasodilatation was almost as potent as that achieved by physiologically relevant concentrations of acetylcholine (10 nM–1 μM; Figure 27.2D). The 4αPDD-induced vasodilatation required a functionally intact endothelium, and endothelial inactivation by mechanical damage prevented vasodilatation to 4αPDD. This indicates that the 4αPDD-induced vasodilatation is strictly endothelium dependent. This was further supported by the observation that 4αPDD just caused vasodilatation when applied to the luminal and thus endothelium-covered face of the artery. Neither vasodilatation nor vasoconstriction occurred when 4αPDD was directly applied to the extravasal side (i.e., to the smooth muscle), thus indicating that TRPV4 is most likely not expressed in the vascular smooth muscle of the carotid artery or is not of considerable importance in smooth muscle cell functions. This is also in line with the lack of TRPV4–mRNA expression as determined by single-cell RT-PCR analysis in freshly isolated smooth muscle cells. That 4αPDD-induced vasodilatation is indeed caused by opening of endothelial TRPV4 is further supported by the observation that 4αPDD elicited vasodilatation

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with a KD of 0.3 μM, which is in fact similar to the KD reported for TRPV4 activation [24]. 4αPDD-induced vasodilatation was prevented by buffering intracellular [Ca2+]i with BAPTA-AM in the endothelium and by the TRPV4 channel blocker ruthenium red (1 μM; Figure 27.2D), indicating that 4αPDD exerts its vasodilating effect by inducing Ca2+ inﬂux and subsequently synthesis of endothelial vasodilators. Regarding the classic acetylcholine-induced vasodilatation, we found that ruthenium red, if intraluminally applied, also reduced this endothelium-dependent vasodilatation, although only modestly. The sole intraluminal application of ruthenium red was without effect on basal vessel diameter. These observations suggest that TRPV4 does not contribute substantially to either agonist-induced Ca2+ signaling and thus vasodilatation or to basal control of vascular tone. This also suggests that ruthenium red does not exert gross unspeciﬁc effects or other effects caused by blocking ryanodine-sensitive Ca2+-release channels in smooth muscle.

TRPV4-MEDIATED VASODILATATION IS NITRIC OXIDE DEPENDENT IN CAROTID ARTERIES Inhibition of nitric oxide synthase alone or in combination with the blockade of prostacyclin synthesis almost completely suppressed the 4αPDD-induced vasodilatation (Figure 27.2E), suggesting that this type of vasodilatation largely relies on the synthesis and action of nitric oxide (NO), whereas the other two major vasodilator systems (i.e., the prostacyclin system or the EDHF system) do not seem to make signiﬁcant contributions to the vasodilating effect of 4αPDD in this conduit artery. NO synthesis following TRPV4 activation is most likely related to the Ca2+ inﬂux through TRPV4 channels and subsequent stimulation of Ca2+-dependent eNOS (Figure 27.1). Regarding EDHF-mediated vasodilatation, Ca2+-dependent activation of endothelial KCa channels of the KCa3.1 and KCa2.3 types and subsequent endothelial hyperpolarization have been considered prerequisites for generating the EDHF signal (Figure 27.1); selective inhibition of these KCa channels abolishes EDHF-type vasodilatation in many vessels [4] and species including the rat carotid artery [6]. Moreover, EDHF-type vasodilatations have been shown to become more important when vessel size decreases [37]. In the large-conduit carotid artery, in which the EDHF system is apparently less important than the NO system, 4αPDD-induced TRPV4 activation and subsequent Ca2+ entry did not cause major EDHF-mediated vasodilatation (Figure 27.2E). In contrast, 4αPDD was able to produce EDHF-mediated vasodilatation in small-sized arteries (A. gracilis 200 μM in diameter). Therefore, pharmacological opening of TRPV4 appears to be sufﬁcient to induce EDHF-type vasodilatation in small-sized arteries, in which EDHF plays a signiﬁcant role.

A FUNCTIONAL ROLE OF TRPV4 IN ENDOTHELIAL MECHANOTRANSDUCTION? In keeping with the proposed mechanosensitivity of TRPV4 [21,25,31,36], we speculated that TRPV4 activation and Ca2+ entry may occur by mechanical stimulation of the endothelium by increased ﬂuid viscosity and thus shear stress. As stated in

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previous paragraphs, shear stress– or ﬂow-induced elevations of endothelial [Ca2+]i are due to both Ca2+ inﬂux and Ca2+ release from internal stores [9,12]. Moreover, such a shear stress–induced increase in [Ca2+]i is prevented by strongly buffering extracellular Ca2+ or by the MSC and TRP blocker Gd3+ [9], indicating that an increase in shear stress activates a “directly” or “indirectly” mechanosensitive Ca2+entry channel. In pressure myography of small vessels, shear stress can be experimentally increased by increasing the viscosity of the perfusion medium by adding dextran. The advantage of this procedure is that pressure gradients are not changed during the experiments, which could lead to additional myogenic effects. By adding 5 percent dextran to the perfusion buffer, the viscosity of the buffer increases from 0.7 to 2.9 mPa*s, which leads to an increase of shear stress from 1 to 3 dyn/cm2 in carotid arteries and from 8 to 19 dyn/cm2 in small-sized arteries, according to the law of Hagen-Poiseuille: τ = 4Q/r3; τ = shear stress; η= viscosity; Q = ﬂow; and r = radius. These experiments revealed that such an increase in shear stress caused substantial vasodilatation of the rat carotid artery (Figure 27.2F), which was strictly NO dependent as an inhibitor of NO-synthesis NG-nitro-L-arginine (L-NNA) completely abolished this vasodilatation. This NO dependency of shear stress–induced vasodilatation is also in agreement with ﬁndings in arteries of humans [38] and other species [39], whereas in mice both prostaglandins as well as NO mediate this type of vasodilatation [40]. Similar to sensitivity of 4αPDD-induced vasodilatation to the pharmacological TRPV4 inhibition, shear stress–induced vasodilatation in the rat carotid artery was greatly blocked by ruthenium red, suggesting TRPV4’s involvement in this response. In addition, the buffering of endothelial [Ca2+]i with BAPTAAM resulted in complete suppression of shear stress–induced vasodilatation, which clearly demonstrates that this is a Ca2+-dependent event and that it mostly depends on Ca2+-dependent NO generation. Inhibition of protein kinase C and of tyrosine kinases was without effect, suggesting that protein phosphorylation in general or a potential TRPV4 phosphorylation by one of these kinases does not seem to play a major role in shear stress–induced vasodilatation. Importantly, shear stress–induced vasodilatation was prevented by inhibition of PLA2 and thus the release of arachidonic acid in rat carotid arteries. Release of AA and production of its metabolites in response to ﬂow is well documented in cultured endothelial cells [41], and a role of AA metabolites in ﬂow-induced vasodilatation has been proposed previously [40]. With respect to TRPV4, exogenously applied AA has been shown to activate rat TRPV4 as shown here and cloned TRPV4 previously [33] and endogenously produced AA mediates mechanical activation (i.e., by cell swelling) [32]. These roles of AA in both shear stress–induced vasodilatation and TRPV4 activation tempt us to speculate that PLA2-mediated release of AA following shear stress stimulation mediates TRPV4 activation. This interpretation also implies that TRPV4 is unlikely to be the mechanosensor per se. Nonetheless, AA-dependent TRPV4 activation might be an essential component in the signal transduction mechanism of endothelial mechanotransduction.

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Collectively, these observations suggest that Ca2+ entry through endothelial TRPV4 channels triggers NO-dependent vasodilatation in the endothelium of the rat CA (conduit artery) and NO- and EDHF-dependent vasodilatation in small-sized A. gracilis (a more resistance artery-like vessel). Moreover, it is tempting to speculate that endothelial TRPV4 channels are involved in endothelial mechanosensing of shear stress–induced vasodilatation. Thus among the numerous TRP channels expressed in the endothelium, TRPV4 might be speciﬁcally assigned to endothelial mechanotransduction.

ENDOTHELIAL TRP CHANNELS AND CARDIOVASCULAR DISEASE Endothelial dysfunction has contributed to several cardiovascular disease states and especially to the defective regulation of vascular tone in hypertension. Moreover, altered functions of endothelial cation channels have been proposed to contribute to this endothelial dysfunction and increased blood pressure. For instance, in rat models of experimental renal insufﬁciency [42], alterations in endothelial cation channel functions have been described for Ca2+-activated K+ channels, which play a crucial role in EDHF-mediated vasodilatation. Accordingly, impaired EDHF signaling and thus endothelial dysfunction was found in this animal model of chronic renal failure. Moreover, genetic manipulation of expression of endothelial Ca2+-activated K+ channels (i.e., the KCa2.3) increases myogenic responsiveness and leads to increased systemic blood pressure. Regarding endothelial Ca2+-permeable cation channels, alterations in mechanosensitive cation channels (MSC) have reported in rat models of genetic hypertension (spontaneously hypertensive rats) and of salt-sensitive hypertension [16,17,43]. Also it is not sufﬁciently investigated whether alterations in endothelial TRP channels also contribute to endothelial dysfunction and increased blood pressure. However, emerging and already existing TRP knockout mice may further elucidate the functional role of speciﬁc TRP in endothelium-dependent control of vascular tone and thus blood pressure control. For instance, the abnormal endothelium-dependent relaxation in TRPC4 knockout mice shows that at least this channel of the canonical TRPC subfamily is important for appropriate endothelial function [23]. Moreover, a recent report on increased myogenic tone and hypertension in TRPC6 knockout mice [44] indicates that also alterations in TRP function in vascular smooth muscle contribute to defective regulation of vascular tone. Whether endothelial functions are disturbed in these mice is currently under investigation. With respect to TRPV4, cardiovascular phenotyping of these mice is superﬁcial so far and thus incomplete [45], and further studies are needed to determine whether endothelial dysfunction and abnormal regulation of vascular tone are present in these mice. Although conventional or even conditional single or double knockout strategies are helpful in elucidating the functional role of the respective TRP channel, one should also consider that early compensation during embryonic development and even rapid compensation in adult animals may occur, thus masking speciﬁc roles of a single channel.

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DO ENDOTHELIAL CA2+-PERMEABLE CATION CHANNELS LIKE TRPV4 REPRESENT NOVEL PHARMACOTHERAPEUTIC TARGETS FOR ANTI-HYPERTENSIVE THERAPY? Endothelial TRP channels and especially TRPV4 can be considered important regulators of vascular tone by modulating intracellular Ca2+ signaling and thus adequate synthesis of vasodilating factors. The functional importance of these ion channels may therefore suggest that they may represent novel pharmacotherapeutic targets in addition to the well-known voltage-gated calcium channels in vascular smooth muscles. From a more general point of view, small molecule openers of TRP channels are more promising than blockers regarding a potential antihypertensive efﬁcacy or therapeutic utility. This based on the assumption that a pharmacological activation of TRP channels and thus an increased Ca2+-inﬂux would improve endothelial function by supporting the synthesis of endothelial vasodilators which is in fact Ca2+dependent. Moreover, it should be considered that a speciﬁc channel is not similarly expressed in vascular smooth muscle since pharmacological activation of this channel would have counteracting vasocontracting effects, thus precluding the usefulness of such a compound. Nonetheless, within the different TRP expressed in the vascular wall, TRPV4 fulﬁlls some of these criteria, and a fairly selective-opener (i.e., 4αPDD) is available to test this idea. As described here, there are some indications that 4αPDD might have blood pressure lowering properties: First, by opening of endothelial TRPV4, 4αPDD would stimulate Ca2+-inﬂux and increase [Ca2+]i, which in turn stimulates the synthesis of vasodilator NO in larger vessel as well as NO- and EDHF-mediated vasodilatation in small resistance-sized arteries and arterioles. Second, 4αPDD exerts vasodilating effects already at submicromolar concentrations and is a completely synthetic compound. This may indicate moderate selectivity. Third, 4αPDD does not induce vasoconstriction in carotid arteries or small-sized arteries as shown here, which is most likely explained by the fact that TRPV4 is not expressed in smooth muscle of these arteries. However, what about potential side effects? It should be considered that TRPV4 is also expressed in epithelia, i.e., airway and kidney epithelia, in the autonomic nervous system, and especially in vessel-innervating sympathetic nerves, and importantly in the ear and in the brain, e.g., in neurosensory cells of the circumventricular organs which are responsive to systemic osmotic pressure. Therefore, it remains to be determined whether TRPV4 openers—as new kind of antihypertensive drugs— might be of clinical usefulness.

REFERENCES [1]

Furchgott, R.F. and Zawadzki, J.V., The obligatory role of endothelial cells in the relaxation of arterial smooth muscle by acetylcholine, Nature, 288, 373–76, 1980. [2] Pohl, U. et al., Crucial role of endothelium in the vasodilator response to increased ﬂow in vivo, Hypertension, 8, 37–44, 1986.

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[7]

[8] [9] [10] [11] [12]

[13] [14]

[15] [16]

[17]

[18] [19]

[20]

[21] [22] [23]

387

Grifﬁth, T.M., Endothelium-dependent smooth muscle hyperpolarization: do gap junctions provide a unifying hypothesis? Br. J. Pharmacol., 141, 881–903, 2004. Busse, R. et al., EDHF: bringing the concepts together, Trends Pharmacol. Sci., 23, 374–380, 2002. Bredt, D.S. and Snyder, S.H., Isolation of nitric oxide synthetase, a calmodulinrequiring enzyme, Proc. Natl. Acad. Sci. USA, 87, 682–685, 1990. Eichler, I. et al., Selective blockade of endothelial Ca2+-activated small- and intermediate-conductance K+-channels suppresses EDHF-mediated vasodilation, Br. J. Pharmacol., 138, 594–601, 2003. Kohler, R. et al., Expression and function of endothelial Ca2+-activated K+ channels in human mesenteric artery: a single-cell reverse transcriptase-polymerase chain reaction and electrophysiological study in situ, Circ. Res., 87, 496–503, 2000. Kohler, M. et al., Small-conductance, calcium-activated potassium channels from mammalian brain, Science, 273, 1709–1714, 1996. Hoyer, J., Kohler, R., and Distler, A., Mechanosensitive Ca2+ oscillations and STOC activation in endothelial cells, Faseb. J., 12, 359–366, 1998. Ishii, T.M. et al., A human intermediate conductance calcium-activated potassium channel, Proc. Natl. Acad. Sci. USA, 94, 11651–11656, 1997. Nilius, B. and Droogmans, G., Ion channels and their functional role in vascular endothelium, Physiol. Rev., 81, 1415–1459, 2001. Helmlinger, G., Berk, B.C., and Nerem, R.M., Pulsatile and steady ﬂow-induced calcium oscillations in single cultured endothelial cells, J. Vasc. Res., 33, 360–369, 1996. Falcone, J.C., Kuo, L., and Meininger, G.A., Endothelial cell calcium increases during ﬂow-induced dilation in isolated arterioles, Am. J. Physiol., 264, H653–H659, 1993. Nollert, M.U., Eskin, S.G., and McIntire, L.V., Shear stress increases inositol trisphosphate levels in human endothelial cells, Biochem. Biophys. Res. Commun., 170, 281–287, 1990. Lansman, J.B., Hallam, T.J., and Rink, T.J., Single stretch-activated ion channels in vascular endothelial cells as mechanotransducers? Nature, 325, 811–813, 1987. Hoyer, J., Kohler, R., and Distler, A., Mechanosensitive cation channels in aortic endothelium of normotensive and hypertensive rats, Hypertension, 30, 112–119, 1997. Hoyer, J. et al., Up-regulation of pressure-activated Ca2+-permeable cation channel in intact vascular endothelium of hypertensive rats, Proc. Natl. Acad. Sci. USA, 93, 11253–11258, 1996. Maroto, R. et al., TRPC1 forms the stretch-activated cation channel in vertebrate cells, Nat. Cell Biol., 7, 179–185, 2005. Liedtke, W. and Kim, C., Functionality of the TRPV subfamily of TRP ion channels: add mechano-TRP and osmo-TRP to the lexicon! Cell Mol. Life Sci., 62, 2985–3001, 2005. Clapham, D.E. et al., International Union of Pharmacology. XLIII. Compendium of voltage-gated ion channels: transient receptor potential channels, Pharmacol. Rev., 55, 591–596, 2003. Nilius, B., Droogmans, G., and Wondergem, R., Transient receptor potential channels in endothelium: solving the calcium entry puzzle? Endothelium, 10, 5–15, 2003. Hofmann, T. et al., Subunit composition of mammalian transient receptor potential channels in living cells, Proc. Natl. Acad. Sci. USA, 99, 7461–7466, 2002. Freichel, M. et al., Lack of an endothelial store-operated Ca2+ current impairs agonistdependent vasorelaxation in TRP4-/- mice, Nat. Cell Biol., 3, 121–127, 2001.

388 [24] [25] [26] [27] [28]

[29]

[30] [31] [32] [33] [34] [35]

[36] [37]

[38] [39]

[40] [41] [42] [43] [44] [45]

TRP Ion Channel Function in Sensory Transduction Watanabe, H. et al., Activation of TRPV4 channels (hVRL-2/mTRP12) by phorbol derivatives, J. Biol. Chem., 277, 13569–13577, 2002. Strotmann, R. et al., OTRPC4, a nonselective cation channel that confers sensitivity to extracellular osmolarity, Nat. Cell Biol., 2, 695–702, 2000. Nilius, B. et al., TRPV4 calcium entry channel: a paradigm for gating diversity, Am. J. Physiol. Cell Physiol., 286, C195–C205, 2004. Liedtke, W. and Friedman, J.M., Abnormal osmotic regulation in trpv4−/− mice, Proc. Natl. Acad. Sci. USA, 100, 13698–13703, 2003. Colbert, H.A., Smith, T.L., and Bargmann, C.I., OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans, J. Neurosci., 17, 8259–8269, 1997. Watanabe, H. et al., Heat-evoked activation of TRPV4 channels in a HEK293 cell expression system and in native mouse aorta endothelial cells, J. Biol. Chem., 277, 47044–47051, 2002. Earley, S. et al., TRPV4 forms a novel Ca2+ signaling complex with ryanodine receptors and BKCa channels, Circ. Res., 97, 1270–1279, 2005. Andrade, Y.N. et al., TRPV4 channel is involved in the coupling of ﬂuid viscosity changes to epithelial ciliary activity, J. Cell Biol., 168, 869–874, 2005. Vriens, J. et al., Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4, Proc. Natl. Acad. Sci. USA, 101, 396–401, 2004. Watanabe, H. et al., Anandamide and arachidonic acid use epoxyeicosatrienoic acids to activate TRPV4 channels, Nature, 424, 434–438, 2003. Vriens, J. et al., Modulation of the Ca2+ permeable cation channel TRPV4 by cytochrome P450 epoxygenases in vascular endothelium, Circ. Res., 97, 908–915, 2005. Gao, X., Wu, L., and O'Neil, R.G., Temperature-modulated diversity of TRPV4 channel gating: activation by physical stresses and phorbol ester derivatives through protein kinase C–dependent and –independent pathways, J. Biol. Chem., 278, 27129–27137, 2003. O'Neil R, G. and Heller, S., The mechanosensitive nature of TRPV channels, Pﬂügers Arch., 451, 193–203, 2005. Shimokawa, H. et al., The importance of the hyperpolarizing mechanism increases as the vessel size decreases in endothelium-dependent relaxations in rat mesenteric circulation, J. Cardiovasc. Pharmacol., 28, 703–711, 1996. Joannides, R. et al., Nitric oxide is responsible for ﬂow-dependent dilatation of human peripheral conduit arteries in vivo, Circulation, 91, 1314–1319, 1995. Holtz, J. et al., Flow-dependent, endothelium-mediated dilation of epicardial coronary arteries in conscious dogs: effects of cyclooxygenase inhibition, J. Cardiovasc. Pharmacol., 6, 1161–1169, 1984. Sun, D. et al., Enhanced release of prostaglandins contributes to ﬂow-induced arteriolar dilation in eNOS knockout mice, Circ. Res., 85, 288–93, 1999. Frangos, J.A. et al., Flow effects on prostacyclin production by cultured human endothelial cells, Science, 227, 1477–1479, 1985. Kohler, R. et al., Impaired EDHF-mediated vasodilation and function of endothelial Ca2+-activated K+ channels in uremic rats, Kidney Int., 67, 2280–2287, 2005. Kohler, R. et al., Impaired function of endothelial pressure-activated cation channel in salt-sensitive genetic hypertension, J. Am. Soc. Nephrol., 12, 1624–1629, 2001. Dietrich, A. et al., Increased vascular smooth muscle contractility in TRPC6−/− mice, Mol. Cell Biol., 25, 6980–6989, 2005. Suzuki, M. et al., Impaired pressure sensation in mice lacking TRPV4, J. Biol. Chem., 278, 22664–22668, 2003.

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A New Insight into the Function of TRPV2 in Circulatory Organs Katsuhiko Muraki Aichi Gakuin University

Munekazu Shigekawa Senri Kinran University

Yuji Imaizumi Nagoya City University

CONTENTS Introduction............................................................................................................389 Expression of TRPV2 in Circulatory Organs ...........................................391 Mechanosensation of TRPV2 in Circulatory Organs ...............................391 Potential Activation Mechanisms of TRPV2 by Mechanical Stimuli...............................................................................394 Conclusions............................................................................................................394 References..............................................................................................................395

INTRODUCTION The TRPV subfamily has had increasing attention since some channels in this group have been shown to be sensitive to a broad range of environmental stimuli, including heat, osmosensitivity, and mechanical stress. In addition, TRPV proteins are widely expressed in a range of cell types in lower and higher organisms. Although some TRPVs were originally found in the sensory system, ubiquitous expression in the whole body suggests that they play important roles in both sensory and nonsensory transduction functions. All mammalian homologues of TRPVs are calciumpermeable channels, with TRPV1–4 characterized as moderately calcium-selective cationic channels (Nilius, Voets et al. 2005; O'Neil and Brown 2003; Benham, Davis et al. 2002). This calcium permeability is physiologically important because Ca2+ has an obligatory role in regulating diverse cellular functions (e.g., fertilization, 389

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FIGURE 28.1 (A) Schematic diagram showing mechanical stimuli to which vascular and cardiac walls are exposed. (B) Potential converging pathways involved in activation of TRPV2 by mechanical stimuli: (a) involvement of ankyrin repeats in mechanoactivation of TRPV2; (b) TRPV2 activation by mechanical stimuli–producing factors; (c) activation of TRPV2 through stimulation of cell-adhesion molecules or associated proteins with TRPV2 by mechanical stimuli.

muscle contraction, exocytosis, and so on). There is increasing evidence that TRPV1–4 are sensitive to physical stimuli such as osmolarity, stretching, and shear stress (Liedtke and Kim 2005; O'Neil and Heller 2005). Whereas TRPV4 appears to be crucial for some relevant forms of cellular mechanosensitivity, the activation of TRPVs by mechanostress has not been fully elucidated for all channels of this group (O'Neil and Heller 2005). Cellular responses to stretch or shear stimuli by blood ﬂow are one of the key elements in muscle tone regulation (Figure 28.1A). In cell-attached and inside-out patch-recording modes, membrane stretch applied through the recording pipette activates nonselective cationic channels in vascular smooth muscles (Kirber et al. 1988; Davis et al. 1992; Ohya et al. 1998; see review: Beech et al. 2004). The unitary conductances range from 8 to 64 pS for monovalent cations, and a cationic channel blocker, Gd3+, is effective to block the channel. In whole-cell recordings, application of longitudinal cell stretch or cell swelling by pressure on the patch pipette or hypotonic bath solution also evokes Ca2+-permeable cationic currents in vascular myocytes (Davis et al. 1992). Additionally, in cardiac atrial and ventricular myocytes, nonselective cationic channels are activated by cell swelling as well as membrane stretch (Clemo and Baumgarten 1997; Zhang et al. 2000; Kamkin et al. 2003). Similar nonselective cationic channels sensitive to mechanical stimuli including shear stress are also identiﬁed in vascular endothelial cells (Lansman et al. 1987; Oike et al. 1994; see review: Nilius and Droogmans 2001). Although extensive studies to identify a molecular candidate of these mechanosensitive channels have

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been performed, information is still limited (Kanzaki, Nagasawa et al. 1999; Gillespie and Walker 2001). Nevertheless, the mechanosensitive nature of the channels seems to be conserved in higher organisms for some TRP channels, and it is likely that TRPC1, TRPC6, TRPV2, TRPM4, TRPA1, TRPP1, and possibly TRPV4 are potential candidates for the mechanosensitive channels in various native organs (Maroto et al. 2005; Welsh et al. 2002; Muraki et al. 2003; Iwata et al. 2003; Earley, Waldron et al. 2004; Corey et al. 2004; Nauli et al. 2003; Liedtke 2005). In this section, we focus on expression, function, and mechanosensitivity of TRPV2 (a member of the vanilloid receptor TRP subfamily) in circulatory organs such as vascular smooth muscles, cardiac muscles, and the endothelium. Heat activation and trafﬁcking mechanisms of TRPV2 and expression of TRPV2 in neuronal organs will be discussed in another section (Caterina et al. 1999; Kanzaki, Zhang et al. 1999; Boels et al. 2001; Barnhill et al. 2004; Benham, Gunthorpe et al. 2003).

EXPRESSION

OF

TRPV2

IN

CIRCULATORY ORGANS

TRPV2 is mainly expressed in a subpopulation of medium to large sensory neurons and is also distributed in the brain and spinal cord (Caterina et al. 1999; Bender et al. 2005). In nonsensory organs including vascular and cardiac myocytes, TRPV2 is distributed as mRNA and protein (Kanzaki, Zhang et al. 1999; Muraki et al. 2003; (Iwata et al. 2003; review: O'Neil and Brown 2003). The mRNA expression of TRPV2 is also detected in human pulmonary (Fantozzi et al. 2003) and umbilical vein endothelial cells (Figure 28.2E). Based on mRNA expression of TRPV2 in mice, it is speculated that TRPV2 is widely expressed in arterial myocytes, which can be inﬂuenced by a broad range of blood pressures (Muraki et al. 2003). Although intracellular localization of the protein was evident, some growth factors localized TRPV2 to the plasma membrane (Kanzaki, Zhang et al. 1999; Boels et al. 2001; Iwata et al. 2003). The translocation of TRPV2 was also observed when myotubes were subjected to a cyclic stretch (Iwata et al. 2003). The mechanisms of this translocation of TRPV2 are not, however, clear because PI3 kinase inhibitors are effective in some cells and not in others. Nevertheless, localization of the TRPV2 protein to the plasma membrane without treatment with growth factors is found in rat adult dorsal root ganglions, cerebral cortex (Liapi and Wood 2005), and mouse arterial myocytes (Muraki et al. 2003). Glycosylation of TRPV2 has been suggested (~95 KDa) (Kanzaki, Zhang et al. 1999), but the apparent molecular mass of TRPV2 in cardiac and aortic smooth muscles is 85–90 KDa (Iwata et al. 2003; Muraki and Imaizumi, unpublished), close to the predicted mass of full-length TRPV2 (~86 KDa).

MECHANOSENSATION

OF

TRPV2

IN

CIRCULATORY ORGANS

In vascular smooth muscles and cardiac myocytes, striking new evidence has emerged for TRPV2’s role (Muraki et al. 2003; Iwata et al. 2003) In mouse aortic mycoytes, cell swelling caused by hypotonic solution activated a nonselective cationic channel current (NSCC) and elevated [Ca2+]i. These responses were not affected by diltiazem, a Ca2+ antagonist, and caffeine, a Ca2+ releaser from the sarcoplasmic

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FIGURE 28.2 Activation of human TRPV2-like channel currents in human umbilical vein endothelial cells (HUVEC). Activation of the channel was monitored as an increase in Ca2+ ﬂuorescence signal in the cell. (A) In the perfusion of a bathing solution with Ca2+ (a linear line) and without Ca2+ (a dotted line); 70 percent hypoosmotic stress (227 mOsm) was applied. (B) The 70 percent hypoosmotic stress-induced Ca2+ ﬂuorescence change was markedly inhibited by 1 and 3 μM ruthenium red (RuR). (C) A slight but substantial increase in Ca2+ ﬂuorescence signal (delta ratio) was evident when 4αPDD was present in the bathing solution. As control experiments, 10 μM ATP (shaded column) and 70 percent hypoosmotic solution (solid column) were applied. (D) A modest (20–46°C, open squares) and high (20–60°C, closed circles) heat-evoked change in Ca2+ ﬂuorescence signal was plotted against temperature in the bathing solution. (E) Distribution of human type TRPV2, TRPV4a, TRPV4b, and GAPDH mRNA transcripts in HUVEC, HEK293, human heart, and human brain using PCR ampliﬁcation (35 cycles).

reticulum, whereas responses were effectively inhibited by ruthenium red, a TRPV blocker. Elevation of [Ca2+]i by the hypotonic stimulation but not activation of NSCC was abolished by the removal of external Ca2+, suggesting that Ca2+ entry through NSCC has an obligatory role in elevation of [Ca2+]i. Although TRPV2 and TRPV4, but not TRPV3, were present in mouse aortas as mRNA, 4αPDD, a potent activator of TRPV4, was not effective to aortic myocytes. Signiﬁcant immunoreactivity to

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mouse TRPV2 protein was detected in mouse aortic, mesenteric, and basilar arterial myocytes. Treatment of mouse aortas with TRPV2 antisense oligonucleotides suppressed hypotonic stimulation–induced activation of NSCC and elevation of [Ca2+]i as well as expression of TRPV2 protein. Although further evidence should be gathered, these results suggest that TRPV2 in vascular myocytes is an essential component of NSCC activated by membrane stretch and cell swelling. On the other hand, it was evident that TRPV2 expression was increased in cultured myotubes prepared from δ-sarcoglycan-deﬁcient BIO14.6 hamster and mdx mice (a mouse model of dystrophin-deﬁcient Duchenne muscular dystrophy). Moreover, a cyclic stretch as well as growth factors translocated TRPV2 to the plasma membrane. In BIO14.6 myotubes, 45Ca2+ uptake was greater compared with the controls and signiﬁcantly inhibited by RuR. Myocyte damage measured by releasing creatine phosphokinase (CK) from myotube cytoplasm was increased in BIO14.6 myotubes to which a cyclic cell stretch was applied. In the presence of RuR, stretchinduced myocyte damage was signiﬁcantly suppressed. Treatment of BIO14.6 myotubes with TRPV2 antisense oligonucleotides reduced the TRPV2 protein expression and elevation of [Ca2+]i and CK release by cyclic cell stretch, strongly indicating that TRPV2 has an obligatory role in cell abnormalities. In transgenic mice overexpressing TRPV2 in cardiac myocytes, prominent anasarca was observed at the age of ~180 days. Furthermore, the mice that died peripartum developed globally enlarged hearts. The morphological changes in the transgenic mice were similar to those observed in damaged cardiac muscles with Ca2+ overload. These results demonstrate a close link between TRPV2 expression levels and myocyte damage/loss of in vivo heart function. In Chinese hamster ovary K1 (CHO-K1) cells transfected with mouse TRPV2 (mTRPV2-CHO), membrane stretch through the recording pipette and cell swelling caused by hypotonic solution activated NSCC, while not in control of CHO-K1. Moreover, stretch of mTRPV2–CHO cultured on an elastic silicone membrane elevated their [Ca2+]i (Muraki et al. 2003). Consistently, 45Ca2+ uptake in mTRPV2– CHO was signiﬁcantly greater than in nontransfected cells, and the uptake was further enhanced by cyclic stretch (Iwata et al. 2003). Taken together, these results provide us with new insight that TRPV2 is a mechanosensitive channel. TRPV4 also appears to display mechnosensitive properties, at least to hypotonic stress and possibly to ﬂuid ﬂow or shear stress. Because TRPV4 has been highly expressed in mouse aortic endothelial cells, TRPV4 is a potential candidate as a mechanosensor to shear stress/ﬂuid ﬂow in vascular endothelial cells (Watanabe, Vriens et al. 2002; Vriens, Watanabe et al. 2004). However, it has been reported that TRPV2 is distributed in vascular endothelial cells (Fantozzi et al. 2003; see also Figure 28.2E), suggesting that TRPV2 also has a functional role in sensing mechanical stimuli in the endothelium. As shown in Figure 28.2A, application of 70 percent hypotonic solution to HUVECS elevated [Ca2+]i in 2.2 mM Ca2+-containing bathing solution, but not in the solution without Ca2+. Hypoosmotic solution–induced elevation of [Ca2+]i was effectively inhibited by 1 and 3 μM RuR, strongly suggesting that TRPV-like channels are involved in the response. Exposure of HUVECs to 4αPDD, a TRPV4 activator, in a concentration range between 0.1 and 3 μM caused a slight increase in [Ca2+]i. In response to a mild heat stimulus (20–46°C) HUVECS exhibited

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a modest rise in [Ca2+]i (0.13±0.01, n = 21, Figure 28.2D, open squares). In contrast, a higher heat stimulus (20–60°C, Figure 28.2D, closed circles) caused a signiﬁcantly larger rise once the temperature exceeded ~55°C. Although further extensive studies are required, TRPV2 as well as TRPV4, both of which are expressed in the vascular endothelium, are potential mechanosensors in the vascular endothelium.

POTENTIAL ACTIVATION MECHANISMS OF TRPV2 BY MECHANICAL STIMULI Our studies and other additional data strongly suggest that TRPV2 may be a mechanosensor in circulatory organs. However, the mechanisms involved in opening TRPV2 by membrane stretch or hypoosmotic cell swelling have not been determined. Moreover, at present, the mechanisms in opening TRPV4 by mechanostress are not elucidated. Deletion of ankyrin repeats, which are present in the N-terminal region of TRPVs, abolished heat activation of TRPV4 (Watanabe, Vriens et al. 2002). Because the ankyrin repeats interact with certain cytoskeletal proteins, this region of TRPV2 might also be important for acceptance of applied mechanical stimuli (Figure 28.1B[a]). In addition, it is possible that cell swelling or membrane stretch produces a certain endogenous ligand that activates TRPV2 (Figure 28.1B[b]). Some fatty acids released by cell stretching modify channel activity. Metabolites of arachidonic acid or diacylglycerol might affect TRPV2 as they do TRPV4 and TRPC6 (Watanabe, Vriens et al. 2003; Slish et al. 2002). A micro-domain composed with channels, lipids, receptors, and/or enzymes could also be an effector of local change of membrane tension by membrane stretch, cell swelling, or shear stress (Figure 28.1B[c]). It has been proposed that adhesion molecules and integrins play important roles in endothelial mechanosensing in addition to their involvement in cell attachment and migration (Shyy and Chien 2002). Also TRPV4 has a tight interaction with ryanodine receptors as well as K+ channels, all of which are involved in regulating vascular tones (Earley, Heppner et al. 2005). Further studies may provide a new line of evidence of a tight interaction of TRPV2 with adhesion molecules, cytoskeletal proteins, or signal transduction units. This is important when compared with the delay of opening of the transduction channels after mechanical stimuli in ﬂy mechanoreceptors (submillisecond response time of mechanogated channels in the ﬂy; Earley, Heppner et al. 2005; review: Gillespie and Walker 2001): those of TRPV2 and TRPV4 after physical stimuli seem to be relatively slow (not determined precisely, at least ~1 second), indicating that these channels are mechanosensitive but not mechanogating (Figure 28.1B[d]).

CONCLUSIONS TRPVs including TRPV2 display multimodal activation by diverse chemical and physical stimuli and function as molecular integrators. The available evidence demonstrates that TRPV2 appears to function as a molecular sensor of heat and mechanical stimuli. In circulatory organs, TRPV2 may regulate blood pressure and cell degeneration under some pathophysiological conditions. Other roles of TRPV2 continue to be explored in an attempt to deﬁne the role of translocation of TRPV2 by growth factors. Furthermore, mechanisms of activation of TRPV2 by mechanical stimuli and interactions of TRPV2 with functional molecules should be deﬁned in the future.

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REFERENCES Barnhill, J.C., A.J. Stokes, et al. (2004). RGA protein associates with a TRPV ion channel during biosynthesis and trafﬁcking. J Cell Biochem 91(4): 808–20. Beech, D.J., K. Muraki, et al. (2004). Non-selective cationic channels of smooth muscle and the mammalian homologues of Drosophila TRP. J Physiol 559(Pt. 3): 685–706. Bender, F.L., Y.S.M. Mederos, et al. (2005). The temperature-sensitive ion channel TRPV2 is endogenously expressed and functional in the primary sensory cell line F-11. Cell Physiol Biochem 15(1–4): 183–94. Benham, C.D., J.B. Davis, et al. (2002). Vanilloid and TRP channels: a family of lipid-gated cation channels. Neuropharmacology 42(7): 873–88. Benham, C.D., M.J. Gunthorpe, et al. (2003). TRPV channels as temperature sensors. Cell Calcium 33(5–6): 479–87. Boels, K., G. Glassmeier, et al. (2001). The neuropeptide head activator induces activation and translocation of the growth-factor-regulated Ca(2+)-permeable channel GRC. J Cell Sci 114(Pt. 20): 3599–606. Caterina, M.J., T.A. Rosen, et al. (1999). A capsaicin-receptor homologue with a high threshold for noxious heat. Nature 398(6726): 436–41. Clemo, H.F. and C.M. Baumgarten (1997). Swelling-activated Gd3+-sensitive cation current and cell volume regulation in rabbit ventricular myocytes. J Gen Physiol 110(3): 297–312. Corey, D.P., J. García-Añoveros, et al. (2004). TRPA1 is a candidate for the mechanosensitive transduction channel of vertebrate hair cells. Nature 432(7018): 723–30. Davis, M.J., J.A. Donovitz, et al. (1992). Stretch-activated single-channel and whole cell currents in vascular smooth muscle cells. Am J Physiol 262(4 Pt. 1): C1083–88. Earley, S., T.J. Heppner, et al. (2005). TRPV4 forms a novel Ca2+ signaling complex with ryanodine receptors and BKCa channels. Circ Res. Earley, S., B.J. Waldron, et al. (2004). Critical role for transient receptor potential channel TRPM4 in myogenic constriction of cerebral arteries. Circ Res 95(9): 922–29. Fantozzi, I., S. Zhang, et al. (2003). Hypoxia increases AP-1 binding activity by enhancing capacitative Ca2+ entry in human pulmonary artery endothelial cells. Am J Physiol Lung Cell Mol Physiol 285(6): L1233–45. Gillespie, P.G. and R.G. Walker (2001). Molecular basis of mechanosensory transduction. Nature 413(6852): 194–202. Iwata, Y., Y. Katanosaka, et al. (2003). A novel mechanism of myocyte degeneration involving the Ca2+-permeable growth factor-regulated channel. J Cell Biol 161(5): 957–67. Kamkin, A., I. Kiseleva, et al. (2003). Characterization of stretch-activated ion currents in isolated atrial myocytes from human hearts. Pﬂügers Arch 446(3): 339–46. Kanzaki, M., M. Nagasawa, et al. (1999). Molecular identiﬁcation of a eukaryotic, stretchactivated nonselective cation channel. Science 285(5429): 882–86. Kanzaki, M., Y.Q. Zhang, et al. (1999). Translocation of a calcium-permeable cation channel induced by insulin-like growth factor-I. Nat Cell Biol 1(3): 165–70. Kirber, M.T., J.V. Walsh, Jr., et al. (1988). Stretch-activated ion channels in smooth muscle: a mechanism for the initiation of stretch-induced contraction. Pﬂügers Arch 412(4): 339–45. Lansman, J.B., T.J. Hallam, et al. (1987). Single stretch-activated ion channels in vascular endothelial cells as mechanotransducers? Nature 325(6107): 811–13. Liapi, A. and J.N. Wood (2005). Extensive co-localization and heteromultimer formation of the vanilloid receptor-like protein TRPV2 and the capsaicin receptor TRPV1 in the adult rat cerebral cortex. Eur J Neurosci 22(4): 825–34.

396

TRP Ion Channel Function in Sensory Transduction

Liedtke, W. (2005). TRPV4 plays an evolutionary conserved role in the transduction of osmotic and mechanical stimuli in live animals. J Physiol 567(Pt. 1): 53–58. Liedtke, W. and C. Kim (2005). Functionality of the TRPV subfamily of TRP ion channels: add mechano-TRP and osmo-TRP to the lexicon! Cell Mol Life Sci 62(24): 2985–3001. Maroto, R., A. Raso, et al. (2005). TRPC1 forms the stretch-activated cation channel in vertebrate cells. Nat Cell Biol 7(2): 179–85. Muraki, K., Y. Iwata, et al. (2003). TRPV2 is a component of osmotically sensitive cation channels in murine aortic myocytes. Circ Res 93(9): 829–38. Nauli, S.M., F.J. Alenghat, et al. (2003). Polycystins 1 and 2 mediate mechanosensation in the primary cilium of kidney cells. Nat Genet 33(2): 129–37. Nilius, B. and G. Droogmans (2001). Ion channels and their functional role in vascular endothelium. Physiol Rev 81(4): 1415–59. Nilius, B., T. Voets, et al. (2005). TRP channels in disease. Sci STKE 2005(295): re8. Ohya, Y., N. Adachi, et al. (1998). Stretch-activated channels in arterial smooth muscle of genetic hypertensive rats. Hypertension 31(1 Pt. 2): 254–58. Oike, M., G. Droogmans, et al. (1994). Mechanosensitive Ca2+ transients in endothelial cells from human umbilical vein. Proc Natl Acad Sci USA 91(8): 2940–44. O'Neil, R.G. and R.C. Brown (2003). The vanilloid receptor family of calcium-permeable channels: molecular integrators of microenvironmental stimuli. News Physiol Sci 18: 226–31. O'Neil R.G. and S. Heller (2005). The mechanosensitive nature of TRPV channels. Pﬂügers Arch 451(1): 193–203. Shyy, J.Y. and S. Chien (2002). Role of integrins in endothelial mechanosensing of shear stress. Circ Res 91(9): 769–75. Slish, D.F., D.G. Welsh, et al. (2002). Diacylglycerol and protein kinase C activate cation channels involved in myogenic tone. Am J Physiol Heart Circ Physiol 283(6): H2196–201. Vriens, J., H. Watanabe, et al. (2004). Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4. Proc Natl Acad Sci USA 101(1): 396–401. Watanabe, H., J. Vriens, et al. (2003). Anandamide and arachidonic acid use epoxyeicosatrienoic acids to activate TRPV4 channels. Nature 424(6947): 434–38. Watanabe, H., J. Vriens, et al. (2002). Heat-evoked activation of TRPV4 channels in a HEK293 cell expression system and in native mouse aorta endothelial cells. J Biol Chem 277(49): 47044–51. Welsh, D.G., A.D. Morielli, et al. (2002). Transient receptor potential channels regulate myogenic tone of resistance arteries. Circ Res 90(3): 248–50. Zhang, Y.H., J.B. Youm, et al. (2000). Stretch-activated and background non-selective cation channels in rat atrial myocytes. J Physiol 523 Pt. 3: 607–19.

29

The Role of TRPV4 in the Kidney David M. Cohen Oregon Health & Science University and the Portland Veterans Affairs Medical Center

CONTENTS Activation of TRPV4.............................................................................................398 Activation of TRPV4 by Thermal Stress ..................................................398 Activation of TRPV4 by Lipids ................................................................398 Mechanosensation and TRPV4 .................................................................398 Activation of TRPV4 by Hypotonicity .....................................................399 Activation of TRPV4 by Other Stimuli ....................................................400 TRPV4 in Cellular Osmoregulation......................................................................400 TRPV4 in Systemic Osmoregulation ....................................................................401 Overview....................................................................................................401 Functional Anatomy of TRPV4 in the Mammalian Kidney ....................403 A Model of TRPV4 Function in Kidney Physiology ...............................404 The Role of TRPV4 in Regulating Water Balance in vivo ......................405 Sodium Balance versus Water Balance ...........................................405 Classiﬁcation of Hypernatremia ......................................................405 Water Balance in TRPV4−/− Mouse Models....................................406 Summary ................................................................................................................409 Acknowledgment ...................................................................................................409 References..............................................................................................................409

An understanding of the role of TRPV4 in mammalian kidney physiology begins in the nematode C. elegans. Worms are repelled by steep osmotic gradients, an adaptive mechanism presumably conserved to minimize the risk of acute changes in cell volume. Worms with mutated copies of a particular gene, while remaining fully motile, are oblivious to potentially harmful osmotic gradients; this gene was dubbed osm-9 [1,2]. The OSM-9 protein bore a striking similarity to the transient receptor potential channel implicated in visual signal transduction in Drosophila [3]—the prototypical member of the now well-recognized TRP superfamily [4]. Several years later, the mammalian orthologue of OSM-9 was identiﬁed via homology screening [5,6] and other approaches [7,8]; it eventually became known as TRPV4 [9]. 397

398

TRP Ion Channel Function in Sensory Transduction

ACTIVATION OF TRPV4 ACTIVATION

OF

TRPV4

BY

THERMAL STRESS

Like most other members of the TRPV subfamily, TRPV4 is temperature sensitive. This phenomenon was observed under conditions of both native [10,11] and heterologous [11–13] expression. The role of TRPV4 in thermosensation in vivo, however, is complex. For example, whereas TRPV4-null mice and control mice exhibit similar latencies (time to withdrawal) in response to gradual temperature elevation on a heated plate [14], the null mice prefer a warmer ﬂoor temperature than their wildtype littermates do, when presented with a choice [15].

ACTIVATION

OF

TRPV4

BY

LIPIDS

Although it is unresponsive to the vanilloid TRPV1 agonists capsaicin and resiniferatoxin [5,6], TRPV4 is activated by a variety of lipids including phorbol 12myristate 13-acetate (PMA) and 4α-phorbol 12,13-didecanoate (4α-PDD) [16]. This activating effect operates independently of protein kinase C. TRPV4 is also activated by the endogenous cannabinoid anandamide [17], much like TRPV1 (reviewed in reference 18). The anandamide effect is independent of the G-protein-coupled cannabinoid receptors [16,19]. Rather, anandamide is likely metabolized to arachidonic acid and then, via the action of cytochrome P450 epoxygenase, is further metabolized to one of several epoxyeicosatrienoic acid (EET) compounds. These lipids, particularly 5’,6’-epoxyeicosatrienoic acid, then activate TRPV4. Whether activation of TRPV4 by these EETs is direct or indirect remains unresolved. Exogenously applied EETs activate TRPV4 [17]. But there are numerous catabolic pathways for the EETs [20]; much as the TRPV4-activating ability of arachidonic acid was traced to its EET metabolites, so might these putative effects of EET be attributable to one of its many metabolites. Some lipids, including arachidonic acid and anandamide, indirectly inﬂuence the properties of ion channels by altering their lipid microenvironments (see, e.g., references 21 and 22). As discussed earlier, C. elegans mutated for the osm-9 gene (analogous to mammalian TRPV4) fail to avoid osmotic gradients [1,2]. Interestingly, this phenotype is reproduced in worms mutated for a gene encoding a fatty acid desaturase enzyme (fat-3) essential for synthesis of polyunsaturated fatty acids [23]. This sensory defect can be rescued through dietary supplementation of either arachidonic acid or eicosapentaenoic acid [23], suggesting a profound impact of polyunsaturated fatty acids on TRPV4 function.

MECHANOSENSATION

AND

TRPV4

Although TRPV4 is a reputed mechanosensory channel (reviewed in reference 24), the relationship between this role and its osmosensory function (see next section) remains unclear. Data thus far suggest that the channel is only indirectly gated by hypotonicity and that either lipid-mediated or kinase-dependent signaling intermediates are required [17,25,26]. Although it was not activated by direct patch suction, heterologously expressed TRPV4 was variably responsive to positive pressure in

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399

the whole-cell conﬁguration [6,27]. In addition, TRPV4 was activated by shear stress from laminar bath ﬂow [13]. In a mechanosensory capacity, it is tempting to speculate that TRPV4 may form part of the primary cilium on kidney tubule epithelial cells; these apical structures sense ﬂow of “proto-urine” in the tubular lumen [28,29]. At least one other TRP family member, polycystin-2, is a constituent of this putative ﬂow-sensing ciliary apparatus. However, immunolocalization studies were inconsistent with this distribution for TRPV4 in the kidney [30]. Interestingly, TRPV4 localized to an analogous ciliary structure in cells lining the oviduct of the female reproductive tract where coexpression with polycystin-2 was also observed [31].

ACTIVATION

OF

TRPV4

BY

HYPOTONICITY

In heterologous expression systems, TRPV4 is sensitive to minute changes in ambient osmolarity, giving rise to a robust calcium entry signal [5–7]. Conﬂicting data have emerged concerning the molecular mechanism through which hypotonicity inﬂuences TRPV4 function. Xu et al. noted that TRPV4—heterologously expressed in HEK293 cells or natively expressed in a murine kidney distal convoluted tubule cell line— rapidly undergoes tyrosine phosphorylation in response to hypotonic stress, and this phenomenon was sensitive to inhibition of SRC-family cytoplasmic tyrosine kinases [26]. Correspondingly, several SRC family kinases physically associated with TRPV4. Overexpression of one of these kinases, Lyn, enhanced both basal and hypotonicity-inducible tyrosine phosphorylation of TRPV4, whereas overexpression of dominant-negative acting Lyn partially blocked the response. Systematic mutation of tyrosine residues in TRPV4 indicated that Tyr-253 was the site of regulated phosphorylation. Mutation of this site to any of a number of different residues prevented TRPV4-dependent calcium entry in response to hypotonic shock without inﬂuencing channel targeting to the cell membrane [26]. The essential nature of this residue was seen in both stable [26] and transient [32] transfection experiments. Consistent with a role in TRPV4 activation, other data support a role for tyrosine phosphorylation in the cell response to anisotonic stress. Tyrosine phosphorylation of a range of proteins is upregulated by hypotonicity [33–35], and pharmacological manipulation of global tyrosine kinase or phosphatase activity inﬂuences the osmoregulatory efﬂux of ions and osmotically active organic solutes [34,36,37]. In addition, a number of SRC family cytoplasmic tyrosine kinases are activated by cell swelling and are instrumental in the physiological response to hypotonic stress [38]. Similarly, the SRC family kinase Fyn regulates the tonicity-responsive transcription factor, TonEBP [39]. Moreover, it was later shown that other TRPV channels as well as members of the TRPC and TRPM families are similarly regulated by SRC family cytoplasmic tyrosine kinases [40–44]. In contrast to these ﬁndings, Nilius and colleagues concluded that SRC family kinase activity and phosphorylation of Tyr-253 are dispensable for activation of TRPV4 by hypotonicity [25]. In addition to a different expression model, these authors employed single-cell fura-2-based calcium assay in adherent cells, in contrast to a populationwide assessment in suspended cells as Xu et al. had used [26]. The Nilius group further demonstrated via inhibitor studies that phospholipase A2 (PLA2) was essential for TRPV4 activation by hypotonic stress but not by the lipid agonist,

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TRP Ion Channel Function in Sensory Transduction

4α-PDD [25]. These data were consistent with earlier ﬁndings that hypotonicity activated phospholipase A2-dependent arachidonic acid release [45–47]. Cytochrome P450 epoxygenase—which functions downstream of phospholipase A2 and is required for TRPV4 activation by anandamide—was also a component of the pathway leading to TRPV4 activation by hypotonicity. As an aside, this enzyme complex is not a universal participant among TRPV4-activating pathways; inhibition of P450 epoxygenase prevented calcium entry in response to hypotonic stress but not in response to heat stress [25]. These authors went on to implicate involvement of another tyrosine residue, Tyr-555, in activating TRPV4 by heat and 4α-PDD but, importantly, not by cell swelling [25].

ACTIVATION

OF

TRPV4

BY

OTHER STIMULI

In addition to responding to heat, lipids, and mechanical stress, TRPV4 might be activated by other physical stimuli. Cilia on the epithelial cells lining the oviduct propel mucus (as well as gametes and embryos) over a wide range of viscosities. When oviduct ciliated cells were superfused in vitro with a viscous medium (supplemented with dextran), an abrupt increase in TRPV4 calcium channel activity was noted [48]. This effect of viscosity was not detectable in naïve HeLa cells, but was recapitulated if the HeLa cells were transfected with TRPV4. Pharmacological inhibition of phospholipase A2 blocked the viscosity-inducible TRPV4-like current in oviduct ciliated cells [48], consistent with a mode of activation for TRPV4 postulated by the Nilius group [25].

TRPV4 IN CELLULAR OSMOREGULATION Recent data indicate that TRPV4 regulates water balance at the cellular level. Cells subjected to a hypotonic milieu abruptly swell as a consequence of water entry. Unregulated water entry, facilitated by membrane expression of water channel proteins, constitutes a two-pronged threat: the ensuing increase in hydrostatic pressure jeopardizes cell integrity, while dilution of the cytoplasm adversely affects macromolecular structure and function. Acutely swollen cells jettison ions and organic solutes to permit water efﬂux in a process known as regulatory volume decrease. This phenomenon has been studied in great detail in a number of in vitro model systems. In most, hypotonic stress is rapidly followed by calcium entry (reviewed in reference 49); this proximal signaling event then triggers efﬂux of ions (particularly K+ and Cl−) and osmotically active organic solutes such as taurine and sorbitol (reviewed in reference 50). Although many of the volume-regulatory effector mechanisms triggered by swelling-induced calcium transients are conserved across model systems, the mechanism of calcium entry itself is highly cell type–speciﬁc. In the model of human airway epithelial cells, as in most models, the regulatory volume decrease following hypotonic stress is calcium dependent [51]. In this model, both the hypotonicitydependent calcium entry and subsequent regulatory volume decrease require TRPV4 [52]. Downregulation of endogenous TRPV4 expression with a TRPV4directed antisense prevented activation of a calcium-dependent potassium channel

The Role of TRPV4 in the Kidney

A

401

LCFSN cells (natively express TRPV4)

B

CHO cells (natively lack TRPV4)

TRPV4 antisense Untransfected cell volume TRPV4 transfected `-globin antisense time

5% 100 sec

5% 100 sec

FIGURE 29.1 Role of TRPV4 in cellular osmoregulation. (A) LCFSN cells, which natively express TRPV4, were transfected with antisense directed against TRPV4 or an irrelevant control antisense (beta-globin). Increase in cell volume (percent) was then monitored as a function of time of exposure to a 30 percent hypotonic solution. Regulatory volume decrease response was abolished when TRPV4 expression was downregulated with the TRPV4-directed antisense morpholino oligonucleotide. (B) CHO cells, which natively lack TRPV4 and fail to exhibit regulatory volume decrease in response to hypotonic cell swelling, were compared with TRPV4-transfected CHO cells in a similarly designed experiment. Following exposure of cells to 33 percent hypotonic medium, only the TRPV4-transfected CHO cells exhibited regulatory volume decrease. Figures were adapted from references 52 (A) and 53 (B).

by hypotonicity and completely prevented restoration of normal cell volume. An irrelevant antisense had no effect (Figure 29.1A). These important data were the ﬁrst to link TRPV4 function to the regulatory volume decrease response itself, and the data imply a pivotal role for TRPV4 in water balance at the cellular level. Becker and colleagues went on to show a similar relationship [53]. Rather than blocking regulatory volume decrease in TRPV4-expressing cells, they investigated the process in the Chinese hamster ovary (CHO) cell line, which lacks TRPV4 expression. Naïve CHO cells failed to undergo regulatory volume decrease following osmotic swelling (Figure 29.1B); heterologous expression of TRPV4, however, was permissive for this restorative change in cell volume [53]. So it appears that, at least in some models, downregulation of TRPV4 expression abolishes regulatory volume decrease, and heterologous expression of the channel is sufﬁcient to confer this adaptive response.

TRPV4 IN SYSTEMIC OSMOREGULATION OVERVIEW A physiological role for osmotically responsive TRPV channels is indisputable in C. elegans; among higher eukaryotes, data are emerging more slowly. Initial speculation was fueled by the attractive localization of TRPV4 to key mechanosensory and osmosensory tissues, including the mechanosensitive neurons of the mammalian inner

402

TRP Ion Channel Function in Sensory Transduction

A

B

B

CNT

CCD

DCT Cortex

Interstitium

PT MD

* OMCD Outer Medulla

TAL

tonicity signal DTL ATL

IMCD

Inner Medulla

TRPV4-negative tubule segment

Solute movement

TRPV4-expressing tubule segment Capillary

Water movement Abundant TRPV4 expression

TRPV4 Moderate TRPV4 expression Absent TRPV4 expression

FIGURE 29.2 TRPV4 in the mammalian kidney. (A) TRPV4 expression along the mouse and rat nephron. Black shading indicates strong TRPV4 expression; gray shading indicates less intense TRPV4 expression. Throughout the course of the collecting duct, expression was stronger in the intercalated cells (e.g., arrowheads). The kidney is divided into the cortex and medulla; the latter consists of outer and inner medulla. The outer medulla is further divided into outer stripe and inner stripe (not labeled). PT, proximal tubule; DTL, descending thin limb of the Loop of Henle; ATL, ascending thin limb of the Loop of Henle; MD, macula densa; DCT, distal convoluted tubule; CNT, connecting tubule; CCD, cortical collecting duct; OMCD, outer medullary collecting duct; IMCD, inner medullary collecting duct. The ellipse (marked with *) denotes the section depicted in panel B. (B) In all kidney tubule segments where TRPV4 is present, it is predominantly expressed on the basolateral membrane. TRPV4 expression is also conﬁned to cell types lacking constitutive apical water permeability (see text). Nonetheless, TRPV4 is well positioned to respond to local cues of tonicity in the interstitium rather than in the tubular lumen. As shown in panel A, TRPV4-expressing tubule segments coexist with TRPV4-negative segments at all levels of the kidney from the cortex through the inner medulla. The TRPV4-negative segments transport solute (ﬁlled arrow) and water (open arrow) from the tubule lumen to the interstitium; from the interstitium, solute and water are reabsorbed via capillaries. TRPV4expressing segments transport primarily solute (although the TRPV4-expressing collecting duct cells abundantly transport water in a nonconstitutive fashion, i.e., only in the presence of the water-retentive hormone AVP). Therefore, although TRPV4 is not in contact with lumenal ﬂuid, it is exposed to the dynamic environment of the interstitium; hence, changes in solute or water absorption from TRPV4-negative segments can generate a “tonicity signal” (curved arrows) that, through basolaterally expressed TRPV4, can inﬂuence solute transport in the downsteam TRPV4-positive tubule segments. It is in these distal tubule segments where “ﬁne-tuning” of tubular lumenal ﬂuid composition is achieved. This crosssection is approximately at the level of the ellipse (marked with an *) in panel A. Modiﬁed from references 30, 32, and 54.

The Role of TRPV4 in the Kidney

403

ear hair cells and the cells of the osmosensing blood–brain barrier-deﬁcient circumventricular nuclei [5,12]. TRPV4’s expression in the latter context suggested a role in systemic rather than purely cellular osmoregulation. Arguably the best way to assess TRPV4 function in vivo is through targeted gene deletion. Two groups have done this, generating TRPV4−/− mice using slightly but perhaps signiﬁcantly dissimilar methodologies. Although both sets of knockout mice differ from their wild-type counterparts, they unexpectedly exhibit dissimilar phenotypes (see below). In order to fully appreciate a role for TRPV4 in kidney regulation of water balance, and to speculate on mechanisms through which this might be achieved, it is instructive to review the functional anatomy of TRPV4 expression in the mammalian kidney.

FUNCTIONAL ANATOMY

OF

TRPV4

IN THE

MAMMALIAN KIDNEY

Four reports of the cloning of TRPV4 independently noted expression of the channel in the kidney [5–8], particularly in the distal convoluted tubule. In a person of average size, the kidneys convert ~150 liters of daily glomerular ﬁltrate (potential urine or “proto-urine”) into 0.5–1.5 liters of urine; in so doing, they also reabsorb nearly all of the ﬁltered water and inorganic ions, as well as glucose, amino acids, and a wide variety of essential plasma constituents [54]. This 100-fold reduction in urine volume achieved along the kidney tubule is vital; in its absence, lethal volume depletion and hypotension would ensue in minutes. Such robust water conservation is achieved via a combination of apical water permeability of speciﬁc kidney tubule segments and a hyperosmotic renal medullary interstitium. Together, these factors establish both a route and a driving force for water reabsorption from the tubule lumen into the systemic circulation. Although no data unequivocally impute a role for TRPV4 in this process, clues have emerged from a detailed investigation of the distribution of TRPV4 expression in the kidney. Glomerular ﬁltrate traverses the hyperosmotic renal medulla via the tight hairpinlike Loop of Henle (Figure 29.2A). The cells lining the “descending” arm of the loop are freely water permeant; the ﬁltrate becomes progressively more concentrated as water traverses the tubule. An abrupt transition occurs at the genu of the loop, where the “descending” limb meets the “ascending” limb: here, apical water permeability abruptly ceases. This architecture “traps” the newly concentrated ﬁltrate in the lumen, preventing its passive dilution by water entry as it exits the hypertonic inner medulla of the kidney. Interestingly, expression of TRPV4 is completely absent along the early parts of the kidney tubule where passive water reabsorption takes place; however, precisely at the transition between tubule water permeability and impermeability (at the hairpin turn at the base of the Loop of Henle), TRPV4 expression abruptly emerges (Figure 29.2A; [30]). TRPV4 expression continues thereafter for the length of the kidney tubule. The solitary exception is the highly specialized tubule segment called the macula densa—the only other tubule segment exhibiting constitutive apical water permeability. Here, as in the more proximal water-permeant tubule segments, TRPV4 expression is absent [30]. Therefore, TRPV4 expression in the kidney is entirely restricted to those tubule segments that lack constitutive apical water permeability and where a transcellular osmotic gradient may be expected to develop.

404

A MODEL

TRP Ion Channel Function in Sensory Transduction OF

TRPV4 FUNCTION

IN

KIDNEY PHYSIOLOGY

Throughout the ascending loop and in the distal nephron, TRPV4 expression occurs predominantly at the basolateral membrane of kidney tubular epithelial cells. It is therefore unlikely that the channel comes into direct contact with the lumenal glomerular ﬁltrate. Rather, TRPV4 is ideally situated to sense changes in osmolarity of the medullary interstitium (Figure 29.2B). Such ﬂuctuations occur in response to a variety of physiological states and pathophysiological conditions, including even a simple change in urine ﬂow rate. Under such conditions, TRPV4 may monitor local interstitial water balance. This balance is inﬂuenced not only by the transcellular reabsorption of solute and water from the tubular lumen into the interstitium, but also by the rate of efﬂux of solute and water from this tissue compartment. This latter step is accomplished by the dense capillary plexus of the cortex, and by the vasa recta penetrating the hyperosmotic kidney medulla; these envelop the kidney tubules and are charged with the herculean task of returning roughly six liters of solute-laden interstitial water to the systemic circulation each hour. Basolaterally expressed TRPV4 is well positioned to sense an imbalance between reabsorption of solute and water from the tubular lumen and efﬂux of these plasma constituents from the medulla via its draining capillaries. The interstitium surrounding the proximal tubule cells (which lack TRPV4 expression) is functionally continuous with the interstitium surrounding the TRPV4expressing cells of the cortical distal nephron; therefore, the absence of TRPV4 in the proximal nephron may serve to insulate this high-capacity reabsorptive system from responding to subtle interstitial cues. The distal nephron, in contrast, responsible for “ﬁne-tuning” salt and water balance, may be better poised to respond to these cues. This architecture, where TRPV4-negative tubule segments accompany TRPV4-expressing segments, exists at all levels of the kidney from the cortex to the inner medulla (Figure 29.2A). For example, an acute change in osmolarity of the outer medullary interstitium could regulate activity of basolaterally expressed TRPV4 in cells of the distal nephron; the resulting intracellular calcium signal could then inﬂuence lumenal or basolateral ion transport in that tubule segment. Cells of the proximal tubule, lacking TRPV4 expression, could not respond to this signal from the interstitium. In the outer medulla, one such signal could be overwhelming of the reabsorptive capacity of the vasa recta, leading to accumulation of excess water or solute in the medullary interstitium; this stimulus in turn would be sensed by TRPV4 on the basolateral side of epithelial cells of the thick ascending limb (Figure 29.2B) and thereby “feed forward” to inﬂuence distal solute transport. Few studies have addressed regulation of TRPV4 abundance or function in the kidney in vivo. Aminoglycoside antibiotics are widely prescribed; unfortunately, drugs of this class are limited by their potential oto- and nephrotoxicity. Kitahara and colleagues noted that administration of kanamycin, a prototypical aminoglycoside, decreased renal expression of TRPV4 [55]. This effect, however, was blunted by the antioxidant dihydroxybenzoate when it was coadministered with the antibiotic. These data are consistent with the ability of antioxidant compounds to protect the inner ear from aminoglycoside-induced cochlear hair cell damage (reviewed in reference 56). Aminoglycoside nephrotoxicity is unusual among the toxic nephropathies in that

The Role of TRPV4 in the Kidney

405

copious urinary losses of water and solute (polyuria) generally ensues [57]. Whether this clinical phenomenon is related to the downregulation of TRPV4 expression in the distal nephron—either as cause or consequence—remains to be explored.

THE ROLE

OF

TRPV4

IN

REGULATING WATER BALANCE

IN VIVO

Sodium Balance versus Water Balance As a putative sensor of tonicity, it is likely that TRPV4 responds to perturbations in systemic water balance rather than systemic sodium balance. Serum sodium concentration is largely independent of total body sodium balance; clinically, these two parameters are readily separable. In humans, the differential diagnosis of hypernatremia (a surrogate for plasma hypertonicity) requires an assessment of total body sodium content and water content. In nearly all circumstances, hypernatremia reﬂects water loss with or without sodium loss, although in rare instances it may follow ingestion or administration of concentrated sodium-containing salts (reviewed in references 58 and 59). Total body sodium balance cannot be assessed quantitatively in humans in clinical practice; rather, it is inferred from a physical examination. Findings consistent with hypovolemia (a deﬁcit in total body sodium content) include increased heart rate and decreased blood pressure. In addition, urinary sodium excretion is quantiﬁed as a crude estimate. That is, abnormally low urinary sodium excretion is a marker for decreased total body sodium content (hypovolemia), whereas a “normal” degree of urinary sodium excretion is consistent with intravascular volume repletion and a physiological level of total body sodium content. Classiﬁcation of Hypernatremia Despite the total body sodium content of the individual, hypernatremia nearly always reﬂects water loss—water loss in the absence of sodium loss in the “euvolemic” hypernatremias and water loss in excess of ongoing sodium loss in the “hypovolemic” hypernatremias. The presence or absence of associated sodium loss, while not impacting the hypernatremia per se, assists in identifying the pathological abnormality [58,59]. Euvolemic hypernatremia is caused primarily by excess urinary water loss (diabetes insipidus) or hypodipsia (decreased thirst). Nonurinary sites of water loss are also possible (e.g., water loss through the gastrointestinal tract or through perspiration). Increased water loss alone, unless overwhelming, rarely produces signiﬁcant hypernatremia because it is limited by AVP action on the kidney-collecting duct (when urinary concentrating ability is preserved), and because it produces a powerful urge to drink. In the absence of an appropriate thirst response or when water is unavailable, hypernatremia can quickly ensue despite maximal urinary water conservation. Correction of euvolemic hypernatremia requires water alone. Hypovolemic hypernatremia, in contrast, results from loss of more than just pure water; “wasting” of sodium in the urine or from other sites (e.g., gut, sweat glands, etc.) is also required. If the sodium loss occurs from a nonrenal site, urinary sodium conservation is seen. Aldosterone, upregulated by the hypovolemia, minimizes urinary sodium excretion. In contrast, if the kidneys are the culprit in the sodium loss,

406

TRP Ion Channel Function in Sensory Transduction

this adaptive response is conspicuously absent. Correction of hypovolemic hypernatremia requires repletion of both water and sodium (albeit more of the former). Water Balance in TRPV4−/− Mouse Models It is helpful to interpret the available data for each of the two independently reported TRPV4 knockout models in light of this clinical paradigm. It is also important to emphasize, as had previously been noted, that some discrepancies between these authors’ ﬁndings might have arisen because of the different ways in which the models were generated [14]. Mizuno et al. [60] compared plasma sodium concentration in TRPV4−/− and wild-type mice. In their relatively small sample (n = 10 per group), they were unable to demonstrate a difference between the mice under basal conditions, although there was a nonsigniﬁcant trend toward higher plasma sodium concentration in the TRPV4−/− mice (140.7 + 0.8 vs. 137.9 + 1.0 mEq/l). This group also reported lower urinary sodium levels for the TRPV4−/− mice (123 + 15 vs. 165 + 21 mEq/l). In addition, they noted an insigniﬁcant but seemingly substantial decrease in systolic blood pressure among the knockout mice (105 + 4 vs. 116 + 5 mmHg). These two latter parameters are perhaps consistent with a modest relative decrease in total body sodium content and effective intravascular volume (hypovolemia); however, because these were not balance studies, the discrepancy in urinary sodium concentration might reﬂect in part the greater dilution of the −/− urine (see below). Therefore, no ﬁrm conclusion can be drawn about the total body sodium (“volume”) status in these mice. With respect to water balance in this study, there was a trend toward higher water loss (urine volumes of 1.8 + 0.5 vs. 1.4 + 0.4 ml/d) and greater water intake (4.5 + 1.0 vs. 4.0 + 0.8 ml/d) among the TRPV4−/− mice, although this did not reach the threshold for statistical signiﬁcance. Urine volume is not a surrogate for urinary water excretion, of course, as the urine may be solute rich or solute poor, and the latter will contribute more to free water clearance. Nonetheless, consistent with these ﬁndings was the relative (and again insigniﬁcant) decrease in urinary creatinine, an indirect index of the degree of urinary concentration. With elevated plasma osmolarity, this constellation of ﬁndings could represent either (1) excess water diuresis driving thirst or (2) osmotic diuresis (e.g., from glycosuria) driving thirst. Discrimination between these scenarios requires an assessment of urine osmolarity; a relatively diluted urine is consistent with the former, whereas an “isosthenuric” urine (i.e., urine with a relatively ﬁxed osmolarity approximating that of plasma) suggests the latter. However, the “basal” urine osmolarities were identical in the +/+ and −/− mice in this model. Unregulated renal water loss (diabetes insipidus), which was likely a factor in the Mizuno model (Figure 29.3), generally reﬂects either a defect in hypothalamic production of the water-retentive peptide arginine vasopressin (AVP) or a defect in the renal response to this hormone. Perhaps consistent with the former, basal levels of serum AVP were slightly (albeit insigniﬁcantly) lower in the −/− mice. Consistent with a nephrogenic picture, however, AVP levels in response to an acute osmotic load were dramatically higher in the −/− mice relative to wild-type mice. The osmotic stimulus in this exercise was supraphysiological; the serum osmolarity was acutely

The Role of TRPV4 in the Kidney

A

407

Water intake

Urinary water loss

*

*

Osm (serum)

Osm (urine)

AVP (serum)

*

*

*

Diabetes insipidus

B

Normal

C

†

†

†

Hypodipsia

FIGURE 29.3 Genesis of a water deﬁcit in mammals. In diabetes insipidus (A), excess urinary water loss (ﬁlled arrow) relative to normal (B) is the primary disorder; in primary hypodipsia (C), decreased water intake is primary. The thickness of the horizontal arrows reﬂects the degree of water movement. In the left half of the ﬁgure, primary disturbances are depicted as ﬁlled arrows and secondary compensatory mechanisms as open arrows. The right half of the ﬁgure depicts the biochemical abnormalities expected in each case, relative to normal, with respect to plasma osmolarity, urine osmolarity, and plasma AVP. Abnormalities noted in the model of Mizuno et al. [60] are marked with an asterisk, and those observed in the model of Liedtke and Friedman [61] are marked with a dagger. Note that Mizuno et al. measured plasma sodium concentration rather than osmolarity; the latter is more instructive but may be confounded by other plasma constituents (see text). Many of the abnormalities attributed here to Mizuno et al. did not achieve statistical signiﬁcance in their relatively small sample and were not highlighted in their publication; nonetheless, their internal consistency is striking. In diabetes insipidus, the plasma AVP level is decreased in central (i.e., neurogenic) diabetes insipidus or increased in the setting of AVP-insensitive kidney tubules (nephrogenic diabetes insipidus). Note that other (i.e., nonrenal) routes of water loss may occur. This diagram is not meant to reﬂect the full spectrum of disorders that may give rise to a water deﬁcit.

increased from 310 to 410 using propylene glycol infusion. But in vitro exposure of brain slices to much more subtle hyperosmotic stimuli resulted in a similarly enhanced release of AVP from TRPV4−/− slices relative to slices from the +/+ brains. In addition, independent of AVP, excessive water diuresis can also be seen with gross abnormalities in plasma calcium or potassium concentrations, but neither were present in this model. In aggregate, the data seem most consistent with diabetes insipidus in the TRPV4−/− mice in the model of Mizuno et al. (Figure 29.3). Of note, this interpretation is based primarily on nonsigniﬁcant data trends; nonetheless, these “trends” are internally consistent and hence credible. In the model of Liedtke and Friedman, plasma osmolarity was measured rather than plasma sodium concentration [61]. In contrast to the study of Mizuno et al., Liedtke found a clear-cut increase in plasma osmolarity among the TRPV4−/− mice (300 vs. 295 mosmol/kgH2O). To “unmask” this difference, however, mice were

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single-housed and deprived of food for 24 hours prior to assessment; in the absence of these reﬁnements, no difference was observed. Whereas assessment of plasma osmolarity rather than plasma sodium appears to be a more direct measurement of this physiological parameter, it is subject to confounding in practice. Speciﬁcally, osmotically “ineffective” substances in the plasma such as urea (i.e., blood urea nitrogen) can contribute to measured osmolarity. However, because urea readily crosses most cell membranes, it is not considered an “effective” osmole in vivo. In contrast, other nonsodium solutes are osmotically effective in vivo (e.g., glucose) and do contribute to plasma osmolarity. Therefore, from a clinical perspective, plasma sodium concentration is frequently a more reliable index of a relative water deﬁcit or excess than is plasma osmolarity; when the latter is used in the evaluation of human disease, simultaneous determination of blood urea nitrogen and glucose is generally obtained. Also of note in this study, mice were deprived of food but access to water was not restricted. Dietary intake is the major source of urinary urea, which contributes osmoles to the urine and can act as an osmotic diuretic. Protein intake also dictates blood urea nitrogen, which, although osmotically ineffective, inﬂuences measured osmolarity of the plasma. Therefore, protein starvation can signiﬁcantly impact plasma and urine osmolarity and urine volume independently of water balance. It is unclear if these may have been factors in their study. With respect to an assessment of volume status (total body sodium content) in the study of Liedtke and Friedman, blood pressure and urinary sodium content were not reported. It is in the realm of water balance where the striking ﬁndings emerge. In concert with an elevated plasma osmolarity, and in marked contrast to the observations of Mizuno et al., Liedtke noted dramatically reduced (~30 percent less) water intake among the TRPV4−/− mice [61]. When water was restricted for 24 hours, the discrepancy in plasma osmolarity was magniﬁed, increasing by a further 6 percent in the −/− mice compared to only 3 percent in wild-type mice. This suggested accelerated water loss in the TRPV4 knockout mice, although data comparing urine volume or osmolarity were not available. In aggregate, these data seem most consistent with hypodipsia (i.e., a primary decrease in thirst or osmosensation; Figure 29.3) and, potentially, an inappropriate renal response to this disorder. The data of Mizuno and colleagues [60] and of Liedtke and Friedman [61] suggest dissimilar disorders of water balance. In general, when evaluating a state of plasma hypertonicity, maneuvers designed to impact water access or water loss are paramount. Water restriction, with serial monitoring of an increment in plasma sodium (and osmolarity); detailed analysis of urine volume, osmolarity, sodium, and potassium content (i.e., in metabolic cages); and assessment of water intake would be optimal. The impact of supplemental AVP, if any, on these parameters could then be assessed. It is possible that TRPV4−/− mice will exhibit partial abnormalities at multiple levels. Tissue-speciﬁc targeting of a TRPV4 knockout to either the distal nephron or the lamina terminalis might permit dissection of these events. In the early and ground-breaking studies of these two groups of investigators, it could not be envisioned beforehand that hypertonicity would ensue in the TRPV4−/− mice; in vitro data with TRPV4 as a sensor of hypotonic stress suggested that genetic absence of this protein might predispose to water excess rather than water loss. But now the

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foundation has been laid for a detailed investigation into the role of TRPV4 in the regulation of systemic water balance under basal and pathological conditions.

SUMMARY TRPV4 is activated by a variety of fatty acid metabolites and by a number of physical stimuli including thermal and osmotic stresses. It is likely that tonicity-dependent activation of TRPV4 requires SRC family kinase–dependent tyrosine phosphorylation. Other data support a role for phospholipase A2 and cytochrome P450 epoxygenase activity and involvement of epoxyeicosatrienoic acids. The architecture of subcellular localization of TRPV4 in the mammalian kidney suggests a role for this channel in responding to interstitial signals related to systemic water balance. The TRPV4-expressing tubule segments may then integrate and respond to these signals by altering lumenal solute reabsorption in a “feed forward” regulatory mechanism. The two TRPV4−/− mouse models generated to date have given rise to conﬂicting data: genetic absence of this sensor of hypotonicity induces a state akin to diabetes insipidus in one model and primary hypodipsia in the other. It is likely that more detailed water (and solute) balance studies will establish whether these models are as dissimilar as they initially appear, and will clarify the role of TRPV4 in regulating systemic water balance.

ACKNOWLEDGMENT This work was supported by the National Institutes of Health, the Department of Veterans Affairs, and the American Heart Association.

REFERENCES 1. Colbert HA, Bargmann CI: Odorant-speciﬁc adaptation pathways generate olfactory plasticity in C. elegans. Neuron 14:803–812, 1995. 2. Colbert HA, Smith TL, Bargmann CI: OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans. J Neurosci 17:8259–8269, 1997. 3. Minke B: Drosophila mutant with a transducer defect. Biophys Struct Mech 3:59–64, 1977. 4. Montell C, Rubin GM: Molecular characterization of the Drosophila trp locus: a putative integral membrane protein required for phototransduction. Neuron 2:1313–1323, 1989. 5. Liedtke W, Choe Y, Marti-Renom MA, Bell AM, Denis CS, Sali A, Hudspeth AJ, Friedman JM, Heller S: Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor. Cell 103:525–535, 2000. 6. Strotmann R, Harteneck C, Nunnenmacher K, Schultz G, Plant TD: OTRPC4, a nonselective cation channel that confers sensitivity to extracellular osmolarity. Nat Cell Biol 2:695–702, 2000. 7. Wissenbach U, Bodding M, Freichel M, Flockerzi V: Trp12, a novel Trp-related protein from kidney. FEBS Lett 485:127–134, 2000.

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8. Delany NS, Hurle M, Facer P, Alnadaf T, Plumpton C, Kinghorn I, See CG, Costigan M, Anand P, Woolf CJ, Crowther D, Sanseau P, Tate SN: Identiﬁcation and characterization of a novel human vanilloid receptor-like protein, VRL-2. Physiol Genomics 4:165–174, 2001. 9. Montell C, Birnbaumer L, Flockerzi V, Bindels RJ, Bruford EA, Caterina MJ, Clapham DE, Harteneck C, Heller S, Julius D, Kojima I, Mori Y, Penner R, Prawitt D, Scharenberg AM, Schultz G, Shimizu N, Zhu MX: A uniﬁed nomenclature for the superfamily of TRP cation channels. Mol Cell 9:229–231, 2002. 10. Chung MK, Lee H, Caterina MJ: Warm temperatures activate TRPV4 in mouse 308 keratinocytes. J Biol Chem 278:32037–32046, 2003. 11. Watanabe H, Vriens J, Suh SH, Benham CD, Droogmans G, Nilius B: Heat-evoked activation of TRPV4 channels in a HEK293 cell expression system and in native mouse aorta endothelial cells. J Biol Chem 277:47044–47051, 2002. 12. Guler AD, Lee H, Iida T, Shimizu I, Tominaga M, Caterina M: Heat-evoked activation of the ion channel, TRPV4. J Neurosci 22:6408–6414, 2002. 13. Gao X, Wu L, O’Neil RG: Temperature-modulated diversity of TRPV4 channel gating: activation by physical stresses and phorbol ester derivatives through protein kinase C–dependent and –independent pathways. J Biol Chem 278:27129–27137, 2003. 14. Todaka H, Taniguchi J, Satoh J, Mizuno A, Suzuki M: Warm temperature-sensitive transient receptor potential vanilloid 4 (TRPV4) plays an essential role in thermal hyperalgesia. J Biol Chem 279:35133–35138, 2004. 15. Lee H, Iida T, Mizuno A, Suzuki M, Caterina MJ: Altered thermal selection behavior in mice lacking transient receptor potential vanilloid 4. J Neurosci 25:1304–1310, 2005. 16. Watanabe H, Davis JB, Smart D, Jerman JC, Smith GD, Hayes P, Vriens J, Cairns W, Wissenbach U, Prenen J, Flockerzi V, Droogmans G, Benham CD, Nilius B: Activation of TRPV4 channels (hVRL-2/mTRP12) by phorbol derivatives. J Biol Chem 277:13569–13577, 2002. 17. Watanabe H, Vriens J, Prenen J, Droogmans G, Voets T, Nilius B: Anandamide and arachidonic acid use epoxyeicosatrienoic acids to activate TRPV4 channels. Nature 424:434–438, 2003. 18. Nilius B, Vriens J, Prenen J, Droogmans G, Voets T: TRPV4 calcium entry channel: a paradigm for gating diversity. Am J Physiol Cell Physiol 286:C195–205, 2004. 19. Piomelli D: The molecular logic of endocannabinoid signalling. Nat Rev Neurosci 4:873–884, 2003. 20. Zeldin DC: Epoxygenase pathways of arachidonic acid metabolism. J Biol Chem 276:36059–36062, 2001. 21. Oliver D, Lien CC, Soom M, Baukrowitz T, Jonas P, Fakler B: Functional conversion between A-type and delayed rectiﬁer K+ channels by membrane lipids. Science 304:265–270, 2004. 22. Hilgemann DW: Biochemistry. Oily barbarians breach ion channel gates. Science 304:223–224, 2004. 23. Kahn-Kirby AH, Dantzker JL, Apicella AJ, Schafer WR, Browse J, Bargmann CI, Watts JL: Speciﬁc polyunsaturated fatty acids drive TRPV-dependent sensory signaling in vivo. Cell 119:889–900, 2004. 24. Kung C: A possible unifying principle for mechanosensation. Nature 436:647–654, 2005. 25. Vriens J, Watanabe H, Janssens A, Droogmans G, Voets T, Nilius B: Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4. Proc Natl Acad Sci USA 101:396–401, 2004.

The Role of TRPV4 in the Kidney

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26. Xu H, Zhao H, Tian W, Yoshida K, Roullet JB, Cohen DM: Regulation of a transient receptor potential (TRP) channel by tyrosine phosphorylation. SRC family kinase– dependent tyrosine phosphorylation of TRPV4 on TYR-253 mediates its response to hypotonic stress. J Biol Chem 278:11520–11527, 2003. 27. Suzuki M, Mizuno A, Kodaira K, Imai M: Impaired pressure sensation in mice lacking TRPV4. J Biol Chem 278:22664–22668, 2003. 28. Praetorius HA, Spring KR: The renal cell primary cilium functions as a ﬂow sensor. Curr Opin Nephrol Hypertens 12:517–520, 2003. 29. Delmas P: Polycystins: from mechanosensation to gene regulation. Cell 118:145–148, 2004. 30. Tian W, Salanova M, Xu H, Lindsley JN, Oyama TT, Anderson S, Bachmann S, Cohen DM: Renal expression of osmotically responsive cation channel TRPV4 is restricted to water-impermeant nephron segments. Am J Physiol Renal Physiol 287:F17–24, 2004. 31. Teilmann SC, Byskov AG, Pedersen PA, Wheatley DN, Pazour GJ, Christensen ST: Localization of transient receptor potential ion channels in primary and motile cilia of the female murine reproductive organs. Mol Reprod Dev 71:444–452, 2005. 32. Cohen D: TRPV4 and the mammalian kidney. Pﬂügers Archiv, 2005. 33. Edashige K, Watanabe Y, Sato EF, Takehara Y, Utsumi K: Reversible priming and protein-tyrosyl phosphorylation in human peripheral neutrophils under hypotonic conditions. Archives of Biochemistry & Biophysics 302:343–347, 1993. 34. Tilly BC, van den Berghe N, Tertoolen LG, Edixhoven MJ, de Jonge HR: Protein tyrosine phosphorylation is involved in osmoregulation of ionic conductances. Journal of Biological Chemistry 268:19919–19922, 1993. 35. Sadoshima J, Qiu ZH, Morgan JP, Izumo S: Tyrosine kinase activation is an immediate and essential step in hypotonic cell swelling–induced ERK activation and c-fos gene expression in cardiac myocytes. Embo J 15:5535–5546, 1996. 36. Sachs JR, Martin DW: The role of ATP in swelling-stimulated K-Cl cotransport in human red cell ghosts. Phosphorylation-dephosphorylation events are not in the signal transduction pathway. J Gen Physiol 102:551–573, 1993. 37. Good DW: Hyperosmolality inhibits bicarbonate absorption in rat medullary thick ascending limb via a protein-tyrosine kinase–dependent pathway. Journal of Biological Chemistry 270:9883–9889, 1995. 38. Cohen DM: SRC family kinases in cell volume regulation. Am J Physiol Cell Physiol 288:C483–493, 2005. 39. Ko BC, Lam AK, Kapus A, Fan L, Chung SK, Chung SS: Fyn and p38 signaling are both required for maximal hypertonic activation of the osmotic response elementbinding protein/tonicity-responsive enhancer-binding protein (OREBP/TonEBP). J Biol Chem 277:46085–46092, 2002. 40. Jiang X, Newell EW, Schlichter LC: Regulation of a TRPM7-like current in rat brain microglia. J Biol Chem 278:42867–42876, 2003. 41. Vazquez G, Wedel BJ, Kawasaki BT, Bird GS, Putney JW, Jr.: Obligatory role of Src kinase in the signaling mechanism for TRPC3 cation channels. J Biol Chem, 2004. 42. Jin X, Morsy N, Winston J, Pasricha PJ, Garrett K, Akbarali HI: Modulation of TRPV1 by nonreceptor tyrosine kinase, c-Src kinase. Am J Physiol Cell Physiol 287:C558–563, 2004. 43. Hisatsune C, Kuroda Y, Nakamura K, Inoue T, Nakamura T, Michikawa T, Mizutani A, Mikoshiba K: Regulation of TRPC6 channel activity by tyrosine phosphorylation. J Biol Chem 279:18887–18894, 2004.

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TRP Ion Channel Function in Sensory Transduction

44. Odell AF, Scott JL, Van Helden DF: Epidermal growth factor induces tyrosine phosphorylation, membrane insertion, and activation of transient receptor potential channel 4. J Biol Chem 280:37974–37987, 2005. 45. Thoroed SM, Lauritzen L, Lambert IH, Hansen HS, Hoffmann EK: Cell swelling activates phospholipase A2 in Ehrlich ascites tumor cells. J Membr Biol 160:47–58, 1997. 46. Basavappa S, Pedersen SF, Jorgensen NK, Ellory JC, Hoffmann EK: Swellinginduced arachidonic acid release via the 85-kDa cPLA2 in human neuroblastoma cells. J Neurophysiol 79:1441–1449, 1998. 47. Pedersen S, Lambert IH, Thoroed SM, Hoffmann EK: Hypotonic cell swelling induces translocation of the alpha isoform of cytosolic phospholipase A2 but not the gamma isoform in Ehrlich ascites tumor cells. Eur J Biochem 267:5531–5539, 2000. 48. Andrade YN, Fernandes J, Vazquez E, Fernandez-Fernandez JM, Arniges M, Sanchez TM, Villalon M, Valverde MA: TRPV4 channel is involved in the coupling of ﬂuid viscosity changes to epithelial ciliary activity. J Cell Biol 168:869–874, 2005. 49. Lang F, Busch GL, Volkl H: The diversity of volume regulatory mechanisms. Cell Physiol Biochem 8:1–45, 1998. 50. Wehner F, Olsen H, Tinel H, Kinne-Saffran E, Kinne RK: Cell volume regulation: osmolytes, osmolyte transport, and signal transduction. Rev Physiol Biochem Pharmacol 148:1–80, 2003. 51. Fernandez-Fernandez JM, Nobles M, Currid A, Vazquez E, Valverde MA: Maxi K+ channel mediates regulatory volume decrease response in a human bronchial epithelial cell line. Am J Physiol Cell Physiol 283:C1705–1714, 2002. 52. Arniges M, Vazquez E, Fernandez-Fernandez JM, Valverde MA: Swelling-activated Ca2+ entry via TRPV4 channel is defective in cystic ﬁbrosis airway epithelia. J Biol Chem 279:54062–54068, 2004. 53. Becker D, Blase C, Bereiter-Hahn J, Jendrach M: TRPV4 exhibits a functional role in cell-volume regulation. J Cell Sci 118:2435–2440, 2005. 54. Cohen D: TRPV4 and hypotonic stress, in Molecular sensors for cardiovascular homeostasis, edited by Wang D, New York, Kluwer Academic Publishers, 2005. 55. Kitahara T, Li HS, Balaban CD: Changes in transient receptor potential cation channel superfamily V (TRPV) mRNA expression in the mouse inner ear ganglia after kanamycin challenge. Hear Res 201:132–144, 2005. 56. Wu WJ, Sha SH, Schacht J: Recent advances in understanding aminoglycoside ototoxicity and its prevention. Audiol Neurootol 7:171–174, 2002. 57. Woolfson R, Hillman K: Causes of acute renal failure, in Comprehensive clinical nephrology, edited by Johnson R, Feehally J, London, Mosby, 2000. 58. Adrogue HJ, Madias NE: Hypernatremia. N Engl J Med 342:1493–1499, 2000. 59. Berl T, Kumar S: Disorders of water metabolism, in Comprehensive clinical nephrology, edited by Johnson R, Feehally J, London, Mosby, 2000. 60. Mizuno A, Matsumoto N, Imai M, Suzuki M: Impaired osmotic sensation in mice lacking TRPV4. Am J Physiol Cell Physiol 285:C96–C101, 2003. 61. Liedtke W, Friedman JM: Abnormal osmotic regulation in trpv4−/− mice. Proc Natl Acad Sci USA 100:13698–13703, 2003.

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The TRPV4 Channel in Ciliated Epithelia Yaniré N. Andrade Jacqueline Fernandes Ivan M. Lorenzo Maite Arniges Miguel A. Valverde Universitat Pompeu Fabra

CONTENTS Abstract ..................................................................................................................413 Cell Calcium Signaling in Epithelia .....................................................................413 TRP Channels in Epithelia ....................................................................................414 The Ciliated Epithelium ........................................................................................414 TRPV4 and Cell Volume Regulation in Ciliated Epithelia ..................................415 TRPV4 and Regulation of Ciliary Beat Frequency ..............................................416 Conclusions............................................................................................................418 Acknowledgment ...................................................................................................418 References..............................................................................................................418

ABSTRACT Epithelial layers, such as those lining the airways or the gut, are normally involved in a barrier function as well as in transporting ions and nutrients. Both activities require the polarized distribution of proteins, particularly transport proteins (e.g., ion channels), in luminal and basolateral membranes. Most of the participating transport proteins have already been identiﬁed, and newcomers are received with expectation. Transient receptor potential (TRP) channels are good examples of novel cation transport proteins whose relevance in epithelial physiology is just starting to emerge. In this chapter we focus on the role of the TRPV4 channel in ciliated epithelia.

CELL CALCIUM SIGNALING IN EPITHELIA Many epithelial functions, including absorption, secretion, volume homeostasis and response to pathogens, are linked to changes in intracellular calcium.1 Ciliated epithelia carry out an additional task, the transport of mucus and trapped particles,2 413

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in which Ca2+ signaling is also a key element of regulation (see below). The calcium signal may originate via the activation of entry pathways at the plasma membrane or the release from intracellular stores. Entry of Ca2+ is mainly driven by a favorable electrochemical gradient and the opening of Ca2+-permeable ion channels. We know the most about voltage-gated Ca2+ channels operating in excitable cells. However, only recently the molecular nature of the channels mediating Ca2+ entry in nonexcitable cells, the family of TRP channels, has been identiﬁed. Here we discuss the role of TRP channels, particularly the TRPV4 channel, in generating the Ca2+ signal involved in the regulation of epithelial cell volume and ciliary activity.

TRP CHANNELS IN EPITHELIA The TRP family of channels contains numerous members (27 to date) that present structural similarities.3 The general topology of a TRP subunit consists of six predicted transmembrane domains (TM) with a putative pore loop between the ﬁfth and sixth TMs and intracellular N- and C-terminal regions of variable length. The N-terminal region contains multiple ankyrin repeats in the TRPC, TRPV, TRPA, and TRPN subfamilies, and the C-terminal region presents a domain with enzymatic activity in some members of the TRPM subfamily.3 Several members of the TRP family of channels have been found in epithelial tissues: airways (TRPC1, 4, 6;4 TRPV2, 45,6), intestine and pancreas (TRPV5–67), female reproductive tract (TRPV4 and TRPP1, 28,9), kidney (TRPM3, 6–7;10,11 TRPV1, 4–6;12–14 TRPP1–215), prostate (TRPM8;16 TRPV5–67), inner ear hair cells (TRPA1;17 TRPV4;18 TRPN119), cornea (TRPC420), and epidermal ciliated cells (TRPN119). Of particular interest to this chapter is the contribution of the TRPV4 channel to the physiology of ciliated epithelia. TRPV4 was ﬁrst identiﬁed as a nonselective cation channel rapidly activated under hypotonic conditions when expressed heterologously (reviewed in reference 21). Later studies showed that TRPV4 channels present multiple activating and regulatory sites that allow them to integrate distinct physical and chemical stimuli from the environment, offering a wide range of possible physiological roles. TRPV4 channels also respond to mechanical stress, heat, acidic pH, endogenous ligands, and synthetic agonists such as 4α-phorbol 12,13-didecanoate (4α-PDD).21 TRPV4 mRNA and protein have been identiﬁed in both native ciliated epithelial cells of female reproductive organs8,9 and in cell lines derived from human ciliated airway cells6 where the channel plays key roles in cell volume homeostasis and regulation of ciliary activity. Novel spliced variants of TRPV4 have also been found in airway epithelial cell lines.22 Some of these variants form functional ion channels while others do not oligomerize and are retained intracellularly.

THE CILIATED EPITHELIUM Many epithelial cells lining the airways and the reproductive tract contain projections called cilia, whose function is, in the case of the airways, brushing away foreign particles, pathogens, and allergens. Failure of this ciliary action contributes to respiratory pathology. Similarly, the fallopian tube (oviduct), connecting the uterus with

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the ovaries, transports oocytes and embryos, an essential function for reproduction that also depends on ciliary activity.23 Motile cilia, such as those present in the airways and oviduct, are membrane-delimited extensions of the cell surface of approximately 5–10 μm in length; the motor is a tubulin-based axoneme. The axoneme of each cilia shows nine outer doublets of microtubules and a single central pair of microtubules (9 + 2 structure) formed by heterodimers of α and β tubulin. An important molecule within the cilium structure is the ATPase dynein, a motor linking and displacing two adjacent external doublets, thereby responsible for the cilium movement.2 In contrast to motile cilia, inmotile cilia appear as one cilium per cell (e.g., kidney primary cilia), contain a 9 + 0 axonemal structure, and lack dynein arms.23 An exception is the nodal cilia that express dynein and exhibit a twirling movement. Airway surface microanatomy has received considerable attention over the last years due to its alteration in diseases such as cystic ﬁbrosis.24 It consists of at least two layers: an upper mucus layer (a gel-like polymer network of high-molecular weight mucins) and a lower periciliary liquid layer (PCL) that protects the epithelial cell surface from the mucus layer. Ciliated and mucus-secreting cells form a structural and functional unit and work as a conveyor-belt system for particle transport. In this analogy, cilia provide the power, while the mucus serves as the belt. Under normal physiological conditions, ciliated cells should be able to cope with modiﬁcations in the composition and volume of both compartments, mucus layer, and PCL. Failure to do so may result in pathology.24

TRPV4 AND CELL VOLUME REGULATION IN CILIATED EPITHELIA Cell volume regulation is an important homeostatic mechanism through which cells are able to maintain optimal cell volume in an environment where the extracellular osmotic pressure differs from the intracellular. Cells exposed to a hypotonic extracellular solution (lower relative solute concentration) will initially swell and then recover their original volume. This process is termed regulatory volume decrease (RVD). Conversely, cells exposed to a hypertonic solution (higher relative solute concentration) will initially shrink but then recover their volume to the original size, a process known as regulatory volume increase (RVI). In the absence of extracellular environment modiﬁcations, epithelial cell metabolism and transport processes can also unbalance the ﬁnely tuned osmotic pressure of the cell through the release or generation of osmolytes.25 Although the ability to maintain a constant volume in the face of osmotic stress is important for all cells in the body, the process assumes particular signiﬁcance in epithelial cells. In airways, the luminal face of the epithelial cells is covered by PCL, which modiﬁes its osmolarity under different situations, becoming hyperosmolar (e.g., during cold or dry ventilation) or hypoosmolar (e.g., breathing fog).26 Similarly, osmolarity of oviductal ﬂuid also changes with the ovarian cycle.27 The RVD response typically involves the coordinated activation of Cl− and K+ channels, which permit the passive loss of electrolytes and osmotically obliged water. Cell volume regulation following hypotonic or isotonic swelling in epithelial cells is normally associated with changes in intracellular Ca2+ concentrations28 and

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subsequent activation of Ca2+-dependent ion channels, most likely Ca2+-dependent K+ channels (references 29 and 30 and references therein), although exceptions exist.31 The source of Ca2+ has been difﬁcult to track down, and it varies depending on the stimulus (discussed in reference 29). Most evidence has suggested increased Ca2+ entry via a stretch- or swelling-activated cation channel and subsequent activation of Ca2+-dependent K+ channels. The cloning of the TRPV4 channel provided a molecular candidate to mediate swelling-activated Ca2+ entry, positioning this channel up in the hierarchy of the signaling cascade triggering RVD. We have demonstrated that TRPV4 is expressed and constitutes the only pathway mediating swelling-activated Ca2+ entry required to achieve full RVD in cell lines derived from ciliated human airway epithelia: CFT1–LCFSN tracheal cells6 and HBE bronchial cells (unpublished data). In both cell lines, Ca2+ entry activates either intermediate-30 or large-conductance29 Ca2+-dependent K+ channels, respectively, therefore coupling Ca2+ entry to effective RVD. Interestingly, swelling-dependent TRPV4 activation is missing in cystic ﬁbrosis human airway epithelia,6 resulting in impaired Ca2+-dependent K+ channel activation and defective RVD.30 The actual mechanism explaining the relationship between a defective CFTR Cl− channel protein and TRPV4 as well as the relevance of a defective RVD to cystic ﬁbrosis pathophysiology is unknown at present.

TRPV4 AND REGULATION OF CILIARY BEAT FREQUENCY It has already been mentioned that mucociliary transport plays a fundamental role in the defense against pathogenic agents and, in the case of the oviduct, in the transport of oocytes and embryos to the uterus.23 The relevance of these processes is revealed by the identiﬁcation of defective mucociliary transport in several human respiratory diseases, including cystic ﬁbrosis, asthma, bronchiectasis, and chronic bronchitis,32 as well as in infertility.23 Alterations in the normal functioning of cilia have also been associated to abnormalities in left–right asymmetry33 and the appearance of cysts in the kidneys and other epithelial organs.15 Motile cilia are responsible for mucociliary transport, and a critical factor in the maintenance of appropriate velocity of mucociliary transport is the ciliary beat frequency (CBF).34 Several intracellular signals have been demonstrated to mediate the changes of CBF in response to different stimuli: cAMP, cGMP, nitric oxide, and Ca2+ (reference 8 and references therein). Among them, the role of Ca2+ in the control of CBF is particularly interesting, as it has been associated with the ciliary response to mechanical stimuli. Mechanically stimulated ciliated cells increase intracellular Ca2+ and CBF, a response that is lost in the absence of extracellular Ca2+.35 The hypothesis that mechanical stimulation might be physiologically initiated by changes in mucus viscosity has been present for quite a while, but the cellular mechanism linking the viscous load exerted by the presence of mucus to the control of CBF has just started to be elucidated. Moreover, according to our recent studies,8 a Ca2+ signal links the response of CBF to another important factor determining mucociliary transport: the mucus viscosity, which may change under normal34,36 and pathological conditions.37

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FIGURE 30.1 Effect of viscous load on the ciliary beat frequency (CBF). The CBF dropped ∼35 percent within the low viscosity range of 2 to 37 cP (2–15 percent dextran solutions), but no further decrease was observed at higher viscosities (37–200 cP; corresponding to 15–30 percent dextran solutions). We related this CBF autoregulatory mechanism to the activity of TRPV4 channels. Under low viscosity the channel shows little or no activity. Increasing viscosity activates TRPV4, triggering the autoregulatory response to maintain a correct CBF. Preventing TRPV4-mediated Ca2+ entry (Ca2+-free or Gd3+-containing dextran solutions) or inhibiting PLA2 (upstream of TRPV4 activation) resulted in marked reduction of CBF (modiﬁed from reference 8).

Mucus-transporting cilia appear to present adaptations that allow them to beat under conditions of varying viscosity, thereby preventing the collapse of mucociliary transport, a process known as autoregulation of CBF.8 CBF decreases with increasing external viscosity until it reaches a plateau (Figure 30.1). However, in the absence of extracellular Ca2+ or in the presence of 100 μM Gd3+, oviductal ciliated cells lose their ability to maintain CBF at the same plateau level when challenged with high viscous solutions, suggesting that Ca2+ entry is an important element in the CBF autoregulation.8 This observation prompted us to search for the molecular nature of the Ca2+ entry pathway, as it was the TRP channel’s obvious candidate. Ciliated epithelia of the oviduct express TRPV48 as well as TRPP1 and 29 (also known as PKD1, a regulator of the channel-forming TRPP2 protein [PKD2]).15 More interestingly, TRPV4 (Figure 30.2) and TRPPs localize to the cilia, and both TRPV4 and the TRPP channel complex respond to mechanical stimuli.15,38 However, two pieces of evidence favor the participation of TRPV4 rather than the TRPP channel: (1) no whole-cell cationic current in response to high viscous loading was observed in oviductal ciliated cells dialysed with an anti-TRPV4 antibody, and (2) mechanical activation of the TRPV4 channel, unlike TRPP, requires phospholipase A2 (PLA2) activation and generation of arachidonic acid (AA).8

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FIGURE 30.2 (Color ﬁgure follows p. 234.) Confocal microscopy localization of TRPV4 to motile cilia in a tissue section of the female hamster oviduct. (Left) Nomarsky image. (Right) Detection of TRPV4 (green) and TOPRO-3 nuclear staining (blue). The antibody used to detect TRPV4 is described in references 6, 8, and 22.

CONCLUSIONS TRPV4 has emerged as a candidate to participate in the coupling of ﬂuid viscosity changes to ciliary beat regulation and in cell volume regulation. TRPV4 activation following hypotonic cell swelling39 and viscous load mechanical stimulation8 requires PLA2-dependent formation of AA. These results suggest that the TRPV4 channel is not the volume and mechanosensor per se but is an important piece of the signaling cascade providing the Ca2+ entry pathway. Therefore, it will be interesting to identify the sensor upstream of TRPV4 activation and whether this sensor is shared by the swelling and mechanical-transducing machinery. Another question to evaluate is the association of TRPV4 with the pathophysiology of ciliated epithelia (airways and reproductive organs). Commentaries have already appeared pointing to TRPV4 as a new molecular target to consider in respiratory pathology, but in relation to its presence in airway smooth muscle and immune cells.40,41 The more recent data on TRPV4 involvement in mucociliary transport8 support this view. Deﬁcient functioning of TRPV4 may result in reduced mucus clearance in the airways and failure to transport oocytes and embryos through the oviduct. We believe that this issue will be the focus of future research efforts.

ACKNOWLEDGMENT Work in the authors’ lab has been funded by the Spanish Ministry of Science and Technology (grant number SAF2003-1240), red HERACLES (Fondo de Investigación Sanitaria), and Generalitat de Catalunya (SGR05-266).

REFERENCES 1. Zhang, M.I. & O’Neil, R.G. The diversity of calcium channels and their regulation in epithelial cells. Adv. Pharmacol. 46, 43, 1999.

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2. Satir, P. & Sleigh, M.A. The physiology of cilia and mucociliary interactions. Annu. Rev. Physiol. 52, 137, 1990. 3. Montell, C. The TRP superfamily of cation channels. Sci. STKE. 2005, re3, 2005. 4. Corteling, R.L. et al. Expression of transient receptor potential C6 and related transient receptor potential family members in human airway smooth muscle and lung tissue. Am. J. Respir. Cell Mol. Biol. 30, 145, 2004. 5. Kowase, T. et al. Immunohistochemical localization of growth factor–regulated channel (GRC) in human tissues. Endocr. J. 49, 349, 2002. 6. Arniges, M. et al. Swelling-activated Ca2+ entry via TRPV4 channel is defective in cystic ﬁbrosis airway epithelia. J. Biol. Chem. 279, 54062, 2004. 7. Nijenhuis, T. et al. (Patho)physiological implications of the novel epithelial Ca2+ channels TRPV5 and TRPV6. Pﬂügers Arch. 446, 401, 2003. 8. Andrade, Y.N. et al. TRPV4 channel is involved in the coupling of ﬂuid viscosity changes to epithelial ciliary activity. J. Cell Biol. 168, 869, 2005. 9. Teilmann, S.C. et al. Localization of transient receptor potential ion channels in primary and motile cilia of the female murine reproductive organs. Mol. Reprod. Dev. 71, 444, 2005. 10. Grimm, C. et al. Molecular and functional characterization of the melastatin-related cation channel TRPM3. J. Biol. Chem. 278, 21493, 2003. 11. Schlingmann, K.P. et al. Hypomagnesemia with secondary hypocalcemia is caused by mutations in TRPM6, a new member of the TRPM gene family. Nat. Genet. 31, 166, 2002. 12. Hoenderop, J.G. et al. Epithelial calcium channels: from identiﬁcation to function and regulation. Pﬂügers Arch. 446, 304, 2003. 13. Tian, W. et al. Regulation of TRPV1 by a novel renally expressed rat TRPV1 splice variant. Am. J. Physiol. Renal Physiol. 290, F117, 2005. 14. Cohen, D.M. TRPV4 and the mammalian kidney. Pﬂügers Arch. 451, 168, 2005. 15. Nauli, S.M. & Zhou, J. Polycystins and mechanosensation in renal and nodal cilia. BIOESSAYS 26, 844, 2004. 16. Tsavaler, L. et al. Trp-p8, a novel prostate-speciﬁc gene, is up-regulated in prostate cancer and other malignancies and shares high homology with transient receptor potential calcium channel proteins. Cancer Res. 61, 3760, 2001. 17. Corey, D.P. et al. TRPA1 is a candidate for the mechanosensitive transduction channel of vertebrate hair cells. Nature 432, 723, 2004. 18. Liedtke, W. et al. Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor. Cell 103, 525, 2000. 19. Shin, J.B. et al. Xenopus TRPN1 (NOMPC) localizes to microtubule-based cilia in epithelial cells, including inner-ear hair cells. Proc. Natl. Acad. Sci. USA 102, 12572, 2005. 20. Yang, H. et al. TRPC4 knockdown suppresses epidermal growth factor–induced storeoperated channel activation and growth in human corneal epithelial cells. J. Biol. Chem. 280, 32230, 2005. 21. Nilius, B. et al. TRPV4 calcium entry channel: a paradigm for gating diversity. Am. J. Physiol. Cell Physiol. 286, C195–C205, 2004. 22. Arniges, M. et al. Human TRPV4 channel splice variants revealed a key role of ankyrin domains in multimerization and trafﬁcking. J. Biol. Chem. 281, 1580, 2005. 23. Afzelius, B.A. Cilia-related diseases. J. Pathol. 204, 470, 2004. 24. Knowles, M.R. & Boucher, R.C. Mucus clearance as a primary innate defense mechanism for mammalian airways. J. Clin. Invest. 109, 571, 2002. 25. Hoffmann, E.K. & Dunham, P.B. Membrane mechanisms and intracellular signalling in cell volume regulation. Int. Rev. Cytol. 161, 173, 1995.

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26. Winters, S.L. & Yeates, D.B. Roles of hydration, sodium, and chloride in regulation of canine mucociliary transport system. J. Appl. Physiol. 83, 1360, 1997. 27. Baltz, J.M. Osmoregulation and cell volume regulation in the preimplantation embryo. Curr. Top. Dev. Biol. 52, 55, 2001. 28. McCarty, N.A. & O’Neil, R.G. Calcium signalling in cell volume regulation. Physiol. Rev. 72, 1037, 1992. 29. Fernandez-Fernandez, J.M. et al. Maxi K+ channel mediates regulatory volume decrease response in a human bronchial epithelial cell line. Am. J. Physiol. Cell Physiol. 283, C1705, 2002. 30. Vazquez, E. et al. Defective regulatory volume decrease in human cystic ﬁbrosis tracheal cells because of altered regulation of intermediate conductance Ca2+dependent potassium channels. Proc. Natl. Acad. Sci. USA 98, 5329, 2001. 31. Lock, H. & Valverde, M.A. Contribution of the IsK (MinK) potassium channel subunit to regulatory volume decrease in murine tracheal epithelial cells. J. Biol. Chem. 275, 34849, 2000. 32. Houtmeyers, E. et al. Regulation of mucociliary clearance in health and disease. Eur. Respir. J. 13, 1177, 1999. 33. McGrath, J. et al. Two populations of node monocilia initiate left–right asymmetry in the mouse. Cell 114, 61, 2003. 34. Puchelle, E. et al. Rheological properties controlling mucociliary frequency and respiratory mucus transport. Biorheology 24, 557, 1987. 35. Sanderson, M.J. & Dirksen, E.R. Mechanosensitivity of cultured ciliated cells from the mammalian respiratory tract: implications for the regulation of mucociliary transport. Proc. Natl. Acad. Sci. USA 83, 7302, 1986. 36. Rutllant, J. et al. Rheological and ultrastructural properties of bovine vaginal ﬂuid obtained at oestrus. J. Anat. 201, 53, 2002. 37. Jayaraman, S. et al. Submucosal gland secretions in airways from cystic ﬁbrosis patients have normal [Na(+)] and pH but elevated viscosity. Proc. Natl. Acad. Sci. USA 98, 8119, 2001. 38. Gao, X. et al. Temperature-modulated diversity of TRPV4 channel gating: activation by physical stresses and phorbol ester derivatives through protein kinase C–dependent and –independent pathways. J. Biol. Chem. 278, 27129, 2003. 39. Vriens, J. et al. Cell swelling, heat, and chemical agonists use distinct pathways for the activation of the cation channel TRPV4. Proc. Natl. Acad. Sci. USA 101, 396, 2004. 40. Li, S. et al. Transient receptor potential (TRP) channels as potential drug targets in respiratory disease. Cell Calcium 33, 551, 2003. 41. Liedtke, W. & Simon, S.A. A possible role for TRPV4 receptors in asthma. Am. J. Physiol Lung Cell Mol. Physiol 287, L269–L271, 2004.

31

Protease-Activated Receptors: Mechanisms by Which Proteases Sensitize TRPV Channels to Induce Neurogenic Inﬂammation and Pain Andrew Grant Silvia Amadesi Nigel W. Bunnett University of California, San Francisco

CONTENTS Introduction............................................................................................................422 PAR Cleavage and Activation ...............................................................................422 Molecular Mechanisms of PAR Activation...............................................422 PAR Activating Proteases ..........................................................................423 Mechanisms of PAR Signal Transduction ............................................................426 PAR Coupling to G Proteins and Downstream Signaling Events............426 Mechanisms of PAR Signal Arrest............................................................427 PARs and Protease Signaling in the Nervous System..........................................427 Expression of PARs in the Nervous System.............................................427 Proteases That Activate PARs in the Nervous System .............................429 PAR Regulation of Neuronal Excitability.................................................429 PAR Sensitization of TRPV Channels ......................................................430 Conclusions and Future Perspectives ....................................................................433 Acknowledgment ...................................................................................................434 References..............................................................................................................434

421

422

TRP Ion Channel Function in Sensory Transduction

INTRODUCTION Proteolytic enzymes comprise approximately 2 percent of the human genome.1 Given their abundance, it is not surprising that proteases have diverse biological functions, ranging from the degradation of proteins in lysosomes to the control of physiological processes such as the coagulation cascade. However, a subset of serine proteases (possessing serine residues within their catalytic sites), which may be soluble in the extracellular ﬂuid or tethered to the plasma membrane, are signaling molecules that can speciﬁcally regulate cells by cleaving protease-activated receptors (PARs), a family of four G-protein-coupled receptors (GPCRs). These serine proteases include members of the coagulation cascade (e.g., thrombin, factor VIIa, and factor Xa), proteases from inﬂammatory cells (e.g., mast cell tryptase, neutrophil cathepsin G), and proteases from epithelial tissues and neurons (e.g., trypsins). They are often generated or released during injury and inﬂammation, and they cleave PARs on multiple cell types, including platelets, endothelial and epithelial cells, myocytes, ﬁbroblasts, and cells of the nervous system. Activated PARs regulate many essential physiological processes, such as hemostasis, inﬂammation, pain, and healing. These proteases and their receptors have been implicated in human disease and are potentially important targets for therapy. Proteases and PARs participate in regulating most organ systems and are the subject of several comprehensive reviews.2,3 Within the central and peripheral nervous systems, proteases and PARs can control neuronal and astrocyte survival, proliferation and morphology, release of neurotransmitters, and the function and activity of ion channels, topics that have also been comprehensively reviewed.4,5 This chapter speciﬁcally concerns the ability of PARs to regulate TRPV channels of sensory neurons and thereby affect neurogenic inﬂammation and pain transmission.6,7

PAR CLEAVAGE AND ACTIVATION MOLECULAR MECHANISMS

OF

PAR ACTIVATION

The four cloned PARs share the typical structural features of GPCRs, with seven transmembrane domains, three intracellular and extracellular loops, an extracellular amino terminus, and an intracellular carboxyl terminus. PAR1 was initially identiﬁed by expressing libraries of RNA from thrombin-responsive cells in Xenopus oocytes, and identifying clones that exhibited thrombin responsiveness.8,9 The discovery of PAR2 was accidental and occurred while screening a mouse genomic library using degenerate primers to the bovine neurokinin 2 receptor.10 The existence of at least one further PAR, PAR3, was inferred from the observation that platelets from PAR1 knockout mice still respond to thrombin.11 Finally, PAR4 was identiﬁed by searching expressed sequence tag libraries for PAR-like sequences.12,13 All four PARs share a general common mechanism of activation by proteases, although there are some subtle differences in activation that depend on the receptor and protease in question (these differences will be addressed below). In general, activating proteases cleave receptors at speciﬁc sites within the extracellular amino terminus to reveal a new amino terminus that acts as a tethered ligand, binding to conserved regions of the second extracellular loop, thereby activating the receptor (Table 31.1, Figure 31.1).

Protease-Activated Receptors

423

TABLE 31.1 A Summary of the Major Known PAR-Activating Proteases, Cleavage Sites, Sequences of Activating Peptides, Signaling Mechanisms, and the Sites of PAR Expression PAR1

PAR2

PAR3

PAR4

Activating proteases

Thrombin Factor Xa Trypsin Activated protein C

Trypsin Tryptase Factor VIIa Factor Xa

Thrombin

Thrombin Trypsin Cathepsin G

Cleavage sites ()

LDPR41 S42FLLRN

SKGR34 S35LIGKV

LPIK38 T39FRGAP

PAPR47 G48YPGQV53

Activating peptides

SFLLRN

SLIGKV

None

GYPGQV

Secondary messengers

Gαq/11 PLCβ IP3+DAG Gα12/13 Rho Rho-K, MLCK Gαi cAMP

Gαq/11 PLCβ IP3+DAG ? PLA2 AA Gαs? PKA

Unknown

Gαq/11 ? PKCδ

Locations

Platelets, endothelium, epithelium, neurons, myocytes

Sensory neurons, endothelium, epithelium, myocytes

Platelets, endothelium, myocytes

Platelets, endothelium, epithelium, myocytes

Support for this mechanism of activation comes from the observation that synthetic activating peptides (APs), as short as six amino acids, that correspond to the tethered ligand domain can directly bind to and activate PAR1, PAR2, and PAR4. Although APs generally have a very low potency compared to proteases, they are valuable reagents for investigating the physiological functions of PARs, avoiding the use of proteases that can have many other mechanisms of action. Moreover, APs have been used as the starting materials for developing speciﬁc and potent antagonists and agonists of certain PARs.14,15 However, the peptide corresponding to the tethered ligand for PAR3 is unable to activate this receptor, for unknown reasons. The activating proteases and peptides, intracellular signals, and in vivo locations of the four PARs are summarized in Table 31.1.

PAR ACTIVATING PROTEASES The great diversity of proteases found within living organisms means that there are many different sources for potential activators of PARs. However, the presence of a particular protease within a system does not mean that it will necessarily be able to initiate a signal. Indeed, the capacity of proteases to activate PARs may depend on the presence of binding domains that tether proteases to receptors, of accessory proteins that tether proteases to the cell surface, and of protease inhibitors that modulate enzyme activity.

424

TRP Ion Channel Function in Sensory Transduction

Mast Cells Epithelium

Circulation

Neurons

INJURY AND INFLAMMATION FVIIa, FXa

Tryptase Trypsin IV Trypsin IV SLIGRL 2 1

PAIN TRANSMMISION TRPV Channels

PAR

7 2 3

CLR/RAMP1

CGRP

Sensory nerve ending

SP

CLR/RAMP1 SP CGRP 6

NK1R Spinal neuron

NK1R 5 4 Arteriolar dilation

Venular permeability Granulocyte infiltration

NEUROGENIC INFLAMMATION

FIGURE 31.1 (Color ﬁgure follows p. 234.) Protease cleavage and activation of PAR2 on primary spinal afferent neurons to cause neurogenic inﬂammation and pain. (1) Injury and inﬂammation trigger the generation and release of proteases from mast cells (tryptase), epithelial cells, and neurons (trypsins), and the circulation (FVIIa, FXa) that can cleave PAR2 at the peripheral projections of primary spinal efferent neurons. (2) Cleavage exposes the tethered ligand domain (SLIGRL in mice and rats), which binds conserved regions of the extracellular loop II to activate the receptor. (3) PAR2 activation increases [Ca2+]i and releases CGRP and SP in peripheral tissues. (4) CGRP interacts with the type 1 CGRP receptor (a heterodimer of calcitonin receptor-like receptor, CLR, and receptor activity modifying protein 1, RAMP1) on arterioles to induce dilation and hyperemia. (5) SP interact with the neurokinin 1 receptor (NK1R) on endothelial cells of postcapillary venules to induce gap formation and plasma extravasation. Together, these induce neurogenic inﬂammation. (6) The same stimuli release SP and CGRP in the spinal cord dorsal horn to induce pain transmission. (7) PAR2 sensitize TRPV1 and TRPV4 channels, which causes hyperalgesia to thermal and mechanical stimuli, respectively.

Certain proteases potently activate PARs only when they are concentrated at the cell surface, either through direct interaction with the receptor or by binding to other membrane proteins. As an example of receptor interaction, the anionbinding site of the coagulation factor thrombin binds to charged domains of PAR1 and PAR3, enabling the potent activation of these receptors.16,17 Higher concentrations of thrombin are required to activate PAR4, which lacks a thrombin-binding site.12,13 As an example of interaction with other membrane proteins, the coagulation factors VIIa and Xa interact with tissue factor, an integral membrane protein that is normally expressed by extravascular cells and expressed by endothelial cells and monocytes during inﬂammation, and thereby can potently activate PAR1 and

Protease-Activated Receptors

425

PAR2.18,19 Activated protein C, an anticoagulant protease that degrades coagulation factors Va and VIIa when released from the cell surface, can activate PAR1 when concentrated at the cell surface by the endothelial protein C receptor.20 Interactions between PARs can also inﬂuence the capacity of proteases to activate receptors. The binding of an enzyme to one receptor can activate a different PAR, as in the case of PAR3 and PAR4, the thrombin receptors on murine platelets, which lack PAR1.21 Thrombin interacts with the charged domains of PAR3, which concentrates thrombin at the cell surface of murine platelets to promote thrombin cleavage and activation of PAR4. In addition, there is evidence for intermolecular signaling of certain PARs. The activating peptide sequence for PAR1 (SFLLRN) can activate PAR2, with reduced potency. When coexpressed in the same cell, the tethered ligand domain of PAR1 can interact with the binding site of PAR2, activating an uncleaved receptor.22 Other proteases can potently activate PARs without being concentrated at the cell surface by interactions with receptors or other membrane proteins. For example, trypsins potently activate PAR2 and PAR4 without interacting with receptors or accessory membrane proteins. There are three trypsinogen genes in humans: protease serine (PRSS) I encodes trypsinogen I; PRSS2 encodes trypsinogen II; and PRSS3 encodes mesotrypsinogen.23,24 Trypsinogen IV is a splice variant of mesotrypsinogen.25 Trypsinogen I and II are highly expressed in the pancreas and are secreted into the intestinal lumen, where they can cleave PAR2 on enterocytes.26 These trypsins may also cleave PAR2 in the pancreas during inﬂammation, when trypsins are prematurely activated.27,28 Trypsinogen IV is expressed by brain neurons and epithelial cells of the intestine, airway, and prostate.25,29 Although the physiological functions of trypsinogen IV and mesotrypsinogen are unknown, they can activate PAR1, PAR2, and PAR429 (Bunnett, unpublished). The observations that trypsinogen IV is resistant to polypeptide inhibitors, such as the pancreatic secretory trypsin inhibitor and soybean trypsin inhibitor, and also degrades such inhibitors24,29–31 implies that trypsinogen IV could show sustained activity in tissues, in contrast to trypsinogen I and II, which are susceptible to endogenous inhibitors and unable to degrade them. Posttranslational modiﬁcations of PARs may also affect the ability of certain proteases to activate receptors. This appears to be the case for tryptase, an enzyme that may play an important role in activating PAR2 in the nervous system.32 Tryptase is found in almost all subsets of human mast cells, where it forms up to 25 percent of total cellular protein.33,34 Tryptase is released from mast cells when they degranulate, and because mast cells are often in close proximity to sensory nerve ﬁbers, both in normal and inﬂamed tissues,35 the activation of neuronal PARs by tryptase may be one mechanism by which mast cells regulate neuronal functions. Compared to trypsinogen, tryptase is a weak activator of PAR2. However, deglycosylation of PAR2 at a residue near the cleave site markedly increases the potency with which tryptase activates this receptor.36,37 The reason for this effect is unknown, although glycosylation may impede the interaction of the tryptase tetramer, with its active sites in the center of a doughnut shape, with the PAR2 cleavage site. Different forms of tryptase could also activate PAR2 with different potencies. In humans there are ﬁve different tryptase genes, and further splice variants are present in human mast cells.

426

TRP Ion Channel Function in Sensory Transduction

Recombinant forms of these tryptases have demonstrated mixed results in PAR2 activation assays, with α and βII tryptase activating PAR2 in a lymphoid cell line,38 while βI tryptase showed no PAR activation.39 In addition to tryptase, cells of the immune system release several other PAR-activating proteases. Leukocytes, particularly neutrophils, release cathepsin G, elastase, and proteinase C, all of which can activate PARs.

MECHANISMS OF PAR SIGNAL TRANSDUCTION PAR COUPLING

TO

G PROTEINS

AND

DOWNSTREAM SIGNALING EVENTS

The unique nature of PAR activation, where a single protease molecule can activate multiple receptors, suggests that differences must exist between their signal transduction pathways and those for other GPCRs. Typical GPCR responses involve producing secondary messenger molecules in proportion to the number of receptors occupied by ligands at any one time. A single protease molecule could eventually cleave and activate every PAR on the surface of a cell, so the intensity of PAR responses is determined by the rate at which the receptors are cleaved, in proportion to the number of protease molecules present.40 The identity of the G proteins through which PARs couple to mediate their cellular effects varies depending on the receptor in question and the cell type, allowing PARs to produce a wide range of physiological responses. The majority of studies of the mechanism of signal transduction by PARs have been carried out with PAR1, which can couple through Gαq/11, GαI, and Gα12/13. In platelets, thrombin activation of PAR1 activates Gαq/11, which in turn activates phospholipase C-β1.41 This enzyme catalyzes the hydrolysis of the membrane phospholipid PIP2, releasing inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (DAG), thus releasing Ca2+ from intracellular stores and activating protein kinase C (PKC). Activation of PAR1 on platelets also activates Gα12/13, coupling the receptor to the Rho family of guaninenucleotide exchange factors that regulate Rho-kinase and myosin light chain kinase, mediating thrombin-induced changes in platelet shape.42 In contrast, IP3 and DAG mediate the platelet aggregation and degranulation seen in response to thrombin.43 The potential of PAR1 to also activate GαI, and the ability of the βγ subunits of the G proteins to themselves couple to other signaling pathways (such as phosphatidylinositol 3-kinase activation), further expands the range of intracellular processes mediated by PAR1. Less is known about the signal transduction mechanisms of the other PARs. Activation of PAR2 induces generation of IP3 and a subsequent release of Ca2+ into the cytosol, in transfected cell lines,44 and in cells that endogenously express PAR2, such as neurons and enterocytes,45,46 suggesting that PAR2 couples to Gαq/11. PAR2-mediated release of arachidonic acid and a subsequent generation of prostaglandins occur in enterocytes and transfected epithelial cells, indicating activation of phospholipase A2, although the coupling mechanism by which this occurs is not identiﬁed.26 In addition, there is evidence of an interaction between PAR2 and TRPV channels requiring activation of protein kinase A (PKA), although it is not clear whether this is a result of Gαs activation and the resultant cAMP

Protease-Activated Receptors

427

generation by adenylyl cyclase or whether it is a downstream component of a separate pathway (Bunnett, in press). PAR agonists also couple to the activation of MAP kinases by a variety of mechanisms. One mechanism by which PAR2 activates ERK1/2 involves the βarrestins, cytosolic proteins that interact with activated GPCRs at the plasma membrane. In addition to their role in receptor desensitization and endocytosis, β-arrestins are molecular scaffolds that form an ERK1/2 “signaling module” at the plasma membrane or in endosomes. Thus, PAR2 agonists stimulate the assembly of a large molecular weight complex containing PAR2, β-arrestins, raf-1, and activated ERK1/2, which forms at the plasma membrane or in early endosomes.47 The function of this module is to retain activated ERK1/2 in the cytosol, where they may regulate cytosolic or membrane targets, including ion channels.

MECHANISMS

OF

PAR SIGNAL ARREST

Proteases activate PARs in an irreversible manner; once exposed, the tethered ligand is always available to interact with the cleaved receptor, which could lead to sustained signaling without the existence of efﬁcient mechanisms to attenuate the response. Transient and permanent mechanisms dampen PAR signaling. The transient mechanism involves phosphorylation of PARs by members of the family of G-protein receptor kinases (GRKs), followed by interaction with β-arrestins, which uncouple PARs from heterotrimeric G proteins to desensitize G-protein signaling. GRK3 mediates agonist-induced phosphorylation of PAR1, because GRK3 overexpression enhances thrombin-stimulated PAR1 phosphorylation and suppresses thrombin signaling.48 The overexpression of GRK3 in transgenic mice also attenuates thrombininduced signaling in vivo.49 In endothelial cells GRK5 mediates PAR1 desensitization; GRK5 expression suppresses thrombin signaling, and a dominant-negative GRK5 mutant prolongs thrombin signals.50 The purpose of GRK-induced phosphorylation of a GPCR is to increase its afﬁnity for β-arrestins to interact with GRKphosphorylated GPCRs and disrupt their association with heterotrimeric G proteins to terminate the signal. PAR1 desensitization is diminished in ﬁbroblasts lacking βarrestins.51 Similarly, a PAR2 mutant that is unable to interact with β-arrestins shows sustained signaling and diminished desensitization.47 Receptor endocytosis followed by trafﬁcking to lysosomes for degradation permanently arrests PAR signaling. Agonists promote endocytosis and lysosomal trafﬁcking of PAR1 and PAR2.52–54 Both receptors internalize by a clathrin-mediated process involving the GTPase dynamin, which mediates detachment of clathrincoated pits.55,56 β-arrestins also mediate endocytosis of many GPCRs by acting as adaptors that link receptors to clathrin. However, β-arrestins do not mediate agonistinduced endocytosis of PAR1, which proceeds normally in β-arrestin-deﬁcient ﬁbroblasts.51 Endocytosis of PAR2 does requires β-arrestins. PAR2 agonists induce rapid translocation of β-arrestins from the cytosol to the plasma membrane, followed by a prolonged colocalization of PAR2 and β-arrestins in early endosomes.47,54 Compared to our understanding of endocytosis, less is known about the molecular mechanisms of post-endocytic sorting of GPCRs to lysosomes. Sorting nexin 1, a membrane-associated protein that interacts with the EGF receptor, is required for

428

TRP Ion Channel Function in Sensory Transduction

lysosomal trafﬁcking of PAR1.57 A large amount of the E3 ligase c-Cbl is necessary to sort PAR2 from early endosomes to lysosomes.58

PARs AND PROTEASE SIGNALING IN THE NERVOUS SYSTEM EXPRESSION

OF

PARS

IN THE

NERVOUS SYSTEM

PARs1–4 are expressed in the brain, spinal cord, and associated ganglia, where they have been detected in both neurons and astrocytes. PAR1 mRNA is present in the rat neocortex, cingulate/retrosplenial cortex, subiculum, hypothalamic and thalamic nuclei, and in layers of the hippocampus, cerebellum, and olfactory bulb, where it is present in neurons and astroglia.59 Immunoreactive PAR1 is found in the human cerebellum, temporal and frontal lobes, and the striatum, where it is prominently detected in astrocytes.60 PAR1 is found in dopaminergic neurons in the substantia nigra, GABA-containing granule cells of the olfactory bulb, and glutaminergic neurons of the anterior thalamus. In rats, PAR1 is expressed in spinal cord motoneurons, preganglionic autonomic neurons, and dorsal root ganglia where it colocalizes with the neuropeptides substance P (SP) and calcitonin gene-related peptide (CGRP).61 PAR2 is also expressed by neurons and glial cells throughout the brain, including the hypothalamus,62,63 and by primary spinal afferent neurons containing SP and CGRP.32 Functional experiments conﬁrm the widespread expression of PARs in astrocytes and neurons. Astrocytes express all four PARs,64 and thrombin increases [Ca2+]i in a glioma cell line and in astrocytes by activating both PAR1 and PAR4.65 Astrocytes also express PAR2 and respond to PAR2 agonists.66 PAR1 and PAR2 agonists also signal to dorsal root ganglia neurons to increase [Ca2+]i, providing functional evidence for expression of these receptors.32,46,61 PARs are also expressed in the peripheral nervous system, notably by enteric neurons. A large proportion of myenteric neurons respond to agonists of PAR1, PAR2, and PAR4.46,67,68 Agonists of these receptors depolarize myenteric neurons and increase their excitability.67,68 Approximately 50–60 percent of AH-type neurons (sensory neurons) and slightly fewer S neurons (inter and motor neurons) respond to PAR2 agonists. The PAR2-responsive S neurons also express nitric oxide (NO) synthase and project axons in an anal direction. These are inhibitory motor neurons, and activation would be expected to suppress motility. Submucosal neurons also express PARs, where activation may affect ﬂuid and electrolyte secretion in the intestine. PAR2 is expressed in submucosal neurons of the guinea pig small intestine, some of which also express vasoactive intestinal polypeptides and are thus secretomotor neurons that play an important role in controlling ﬂuid and electrolyte transport.69 PAR2 agonists evoke transient depolarization of submucosal neurons followed by a prolonged hyperexcitability that can last for several hours. This long-term hyperexcitability mimics that observed after degranulation of mast cells, suggesting that mast cell proteases such as tryptase may induce long-term excitability of submucosal neurons through PAR2 activation, which could result in enhanced ﬂuid and electrolyte secretion from the mucosa.70,71

Protease-Activated Receptors

PROTEASES THAT ACTIVATE PARS

429 IN THE

NERVOUS SYSTEM

Proteases from a variety of sources (including the circulatory system, inﬂammatory cells, and even the neurons themselves) have the potential to activate PARs in the central and peripheral nervous systems (Table 31.1). Although many of these enzymes activate PARs on neurons both in vitro and in vivo, in general the proteases that activate neuronal PARs under physiological and pathophysiological conditions have not been unequivocally identiﬁed. Injury and inﬂammation generate several proteases of the coagulation cascade that can activate multiple PARs. Disruption of the integrity of the blood–brain barrier during pathological events such as head trauma and stroke may allow coagulation proteases to enter neural tissue, where they could activate PARs on neurons and astrocytes. During injury these proteases may also activate PARs in peripheral neurons, although this possibility has not been examined. Many inﬂammatory cells are recruited to sites of injury and inﬂammation, where they could release multiple proteases that activate neuronal PARs. In this regard, mast cells may be of particular importance because they are closely associated with sensory nerve ﬁbers in peripheral tissues that contain SP and that are thus nociceptive neurons35 (Figure 31.1). Indeed, tryptase released from mast cells can cleave and activate PAR2 on primary spinal afferent neurons to trigger the release of SP and CGRP in peripheral tissues, where they cause neurogenic inﬂammation, and within the dorsal horn of the spinal cord, where they cause pain transmission.6,27,28,32,72 This mechanism could explain the poorly understood pain of irritable bowel syndrome, where there is a marked inﬂux of tryptase-containing mast cells in close proximity to sensory nerve ﬁbers in the colon, which is accompanied by hyperalgesia to distension of the bowel.73 In addition, neuronal tissue itself expresses proteases that may activate PARs. In the olfactory bulb, mRNA coding for prothrombin and PAR1 colocalize in the same cell populations, suggesting that neuronally derived thrombin may activate neurons by cleaving PAR1.59 Moreover, several trypsinogens, including trypsinogen IV, are also expressed by neural tissues and, if released, could activate PARs.25,29,74 However, nothing is known about the mechanisms that control the release of these neuronal proteases, thus their physiological role remains to be deﬁned.

PAR REGULATION

OF

NEURONAL EXCITABILITY

Accumulating evidence indicates that certain proteases can regulate the excitability of neurons, with important functional consequences. For example, trypsin and tryptase increase the excitability of primary spinal afferent neurons to induce hyperalgesia to thermal and mechanical stimuli6,72 and also to induce hyperexcitability of enteric neurons, with consequences for intestinal secretion and motility.67–69 This increased activity of neurons depends on protease regulation of the function of ion channels. Proteases could regulate ion channels by two general mechanisms: a direct mechanism by which proteases cleave the channel protein itself, and an indirect mechanism by which proteases cleave and activate PARs, which couple to signaling cascades that regulate the activity, localization, or level of expression of the channel.

430

TRP Ion Channel Function in Sensory Transduction

There is good evidence that certain proteases interact directly with ion channels to alter their activity, affecting both ionic transport and electrical excitability of target cells. Channel-activating proteases (CAPs) cleave certain epithelial sodium channels (ENaCs) to increase channel open probability sixfold, an activity mimicked by trypsin, although the precise target of enzymatic cleavage remains obscure.75 Proteases can also regulate the acid-sensing ion channels (ASICs), members of the ENaC family that are found on neurons and that open in response to decreasing extracellular pH.76 ASICs are expressed by nociceptive ﬁbers and may transduce pain signals in response to local acidity.77 ASIC1a and ASIC1b are cleaved by several serine proteases (e.g., trypsin, which only affects ASIC1a, chymotrypsin, and proteinase K), shifting the pH of channel opening and steady-state inactivation to a more acidic pH.78 Proteases can also modify the activity of another neuronal ion channel, the NMDA receptor for the transmitter glutamate. Tissue-plasminogen activator, which is released by cortical neurons upon depolarization, interacts with NMDA receptors on the same neurons to increase their sensitivity to glutamate, leading to an increase in excitotoxic cell death.79 The altered activity occurs as a result of tissue-plasminogen activator cleavage of the NR1 subunit of the NMDA receptor. However, although there is an increasing amount of evidence that proteases can regulate the activity of sensory ion channels, cleavage of TRPV channels by proteases has not been identiﬁed to date. The major known mechanism by which proteases regulate TRPV channels on sensory nerves is through the activation of PARs.

PAR SENSITIZATION

OF

TRPV CHANNELS

Proteases that derive from mast cells (e.g., tryptase) and epithelial tissues (e.g., trypsins) can cleave PAR2 on the ﬁbers of primary spinal afferent neurons to trigger the release of SP and CGRP in peripheral tissues and in the dorsal horn of the spinal cord.6,27,28,32,72 In a similar manner, thrombin activates PAR1 on sensory nerves in peripheral tissues to cause neurogenic inﬂammation.61 However, in contrast to PAR2 agonists, thrombin causes analgesia by mechanisms that are not fully understood.80 The important roles played by TRPV1 and TRPV4 in detecting noxious thermal and mechanical stimuli, respectively, suggests that PAR2-induced thermal and mechanical hyperalgesia may be mediated by sensitization of these ion channels. This sensitization could occur through increased channel activity, decreased channel inhibition, insertion of additional channels into the cell membrane, or a combination of the three. Indeed, agonists of both GPCRs and receptor tyrosine kinases can sensitize TRPV1 by these mechanisms and thereby induce thermal hyperalgesia and inﬂammatory pain. Exposure of sensory neurons to other inﬂammatory mediators (e.g., ATP, bradykinin, NGF), which are released in response to tissue damage or other stresses, induces thermal nociception by sensitizing TRPV1. The mechanisms of this sensitization have been thoroughly investigated. ATP and bradykinin, acting via metabotropic P2Y1 receptors and B2 receptors, respectively, activate PLCβ and consequently PKCε.81,82 Activated PKCε translocates to the plasma membrane and phosphorylates TRPV1 at S502 and S800 to enhance its open probability and inhibit

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FIGURE 31.2 Potential mechanisms by which PAR2 sensitizes TRPV1. (1) Activated PAR2 couples to Gαq11 and activates PLCβ. (2) This pathway leads to the activation of PKCε, which translocates to the plasma membrane. (3) PKCε can phosphorylate TRPV1 at the membrane, resulting in increased open probability and diminished desensitization. (4) PKCε may also activate other downstream kinase, such as PKD, which also phosphorylates and sensitizes TRPV1. (5) In addition, PLCβ hydrolyzes PIP2, which may relieve a tonic inhibition of TRPV1, resulting in channel sensitization. (6) PAR2 agonists also activate PKA, possibly by coupling to Gαs, which also translocates to the plasma membrane where it can phosphorylate and sensitize TRPV1.

its desensitization.83,84 Binding of the membrane phospholipid PIP2 to TRPV1 has been suggested to inhibit channel gating,85 and exposure to NGF sensitizes TRPV1 by removing this inhibition.86 NGF is thought to bind to its tyrosine kinase receptor TrkA to activate PLCγ, which hydrolyzes PIP2 and removes its inhibitory effects on TRPV1. However, a recent study disputes this mechanism of NGF sensitization of TRPV1, suggesting instead that the most signiﬁcant effect occurs through activated TrkA receptors that insert TRPV1-containing vesicles into the plasma membrane.87 Activated TrkA phosphorylates phosphatidylinositol 3-kinase, with a downstream activation of Src kinase leading to phosphorylation of vesicular TRPV1 molecules on Y200, targeting the vesicles to the membrane. Several observations suggest that PAR2 may regulate TRPV1 channels in sensory neurons to induce hyperalgesia to thermal stimuli (Figures 31.1 and 31.2). PAR2 is coexpressed in primary sensory neurons, alongside CGRP, SP, and TRPV1, which are all important components of nociceptive pathways.6,32 Activation of PARs induces neurogenic inﬂammation and nociception and, at subinﬂammatory doses, both mechanical and thermal hyperalgesia.6,32,72 Furthermore, other inﬂammatory agents, such as bradykinin, ATP, and NGF, acting through protein kinases, can sensitize TRPV1.81,82,86 The ﬁrst report of an interaction between PAR2 and TRPV1 was that PAR2 agonists enhance both capsaicin-induced release of CGRP from dorsal root ganglia and fos expression in spinal neurons.27 Capsazepine, a TRPV1 antagonist, also inhibits the hyperalgesia88 and gastric mucus secretion89 induced by PAR2 activation in vivo, providing further evidence for TRPV1’s role in PAR2 regulation of neuronal function. PAR2 agonists also sensitize TRPV1-mediated calcium mobilization and TRPV1 currents in HEK cells expressing TRPV1 and in primary spinal

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afferent neurons.6,7 PAR2-induced thermal hyperalgesia, measured as a decrease in the paw withdrawal latency to a radiant heat, is abolished in mice lacking TRPV1.6,7 Furthermore, nonalgesic doses of PAR2 agonists and capsaicin act synergistically to produce hyperalgesia.6 Together, these results suggest that proteases released during trauma or inﬂammation can potentially activate PAR2 expressed in primary sensory neurons to sensitize TRPV1, resulting in exacerbated responses to TRPV1 activators, exempliﬁed by thermal hyperalgesia. However, there are certain gaps in our understanding of this process. One deﬁciency is that the signal transduction mechanism by which PAR2 sensitizes TRPV1 is not fully understood. PAR2 couples to Gαq/11 and activates PLCβ, releasing IP3 and DAG. PAR2-induced activation of PLCβ could enhance TRPV1 by reducing the levels of its substrate, the membrane phospholipid PIP2, consequently freeing TRPV1 from PIP2-mediated inhibition85,86 (Figure 31.2). In addition, PLCβ-induced release of IP3 and DAG could activate PKC, which phosphorylates TRPV1 to increase its open probability and decrease desensitization.83,84 Indeed, PAR2-mediated potentiation of TRPV1 responses is diminished by PLC inhibitors and also by the nonselective inhibition of PKC inhibitors, supporting both hypotheses.6 The identity of the PKC isozyme responsible for PAR2 sensitization of TRPV1 remains to be determined. One particularly promising candidate is PKCε, which plays a major role in transmitting mechanical and thermal hyperalgesia.90 Also, PKCε phosphorylates TRPV182 to mediate bradykinin-induced sensitization of TRPV1 currents.81 Indeed, selective antagonism of PKCε suppresses PAR2-induced sensitization of TRPV1 currents in nociceptive neurons.7 However, the contribution of other PKC isoforms cannot be excluded, because Gö6976, an inhibitor of conventional PKC isoforms (α, β1, β2, and γ), but not ε, also inhibits PAR2-mediated potentiation of TRPV1 signals.6 This implicates protein kinase D1 (PKD1), as Gö6976 is also a PKD1 inhibitor. Initially thought to be an atypical member of the PKC family (PKCμ), PKD1 is now recognized as a member of a new family of three PKD isozymes that resemble the CAM kinase family.91 Certain novel PKC isoforms activate PKD (e.g., thrombin activation of PKD in platelets involves PKCδ92), which may mediate some of the effects of PKC activation. This possibility is particularly relevant in light of the ﬁnding that PKD1 binds and phosphorylates TRPV1 on S116, enhancing TRPV1-mediated responses, while antisense oligonucleotides directed against PKD1 decreased TRPV1 responses.93 There is also evidence that sensitization of TRPV1 may also occur through activation of PKA. PKA is required for injury-induced hyperalgesia,94 and it phosphorylates TRPV1 on S116 to regulate its desensitization,95,96 thus sensitizing TRPV1 currents in nociceptive neurons.97,98 Another gap in our understanding of the role of PARs in pain transmission is the mechanism of PAR2-induced mechanical hyperalgesia, which is evident after both somatic and visceral mechanical stimuli.72,99 Although the channel that mediates responses to mechanical stimuli is not unequivocally identiﬁed, TRPV4 is a strong candidate. TRPV4 is the mammalian homologue of the C. elegans gene osm-9, which encodes a protein conferring sensitivity to osmotic pressure, touch, and odorants.100 Originally called VR-OAC, this channel responds to small reductions in tonicity; in rats it is expressed in circumventricular organs, which detect changes

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in systemic osmolarity, and in neurosensory structures such as Merkel cells and sensory neurons.101 Based on its structure, ionic selectivity (preference for Ca2+), and thermal sensitivity (>27°C), VR-OAC was placed in the same TRPV family as TRPV4.102,103 The phorbol ester 4α-phorbol 12,13-didecanoate (4αPDD) is a synthetic TRPV4 agonist,104 and the cytochrome P450 product of arachidonic acid, 5’,6’-epoxyeicosatrienoic acid (5’,6’-EET), is a potential endogenous TRPV4 agonist105 (hypoosmotic stimuli leads to cell swelling, phospholipase A2 activation, and release of arachidonic acid106). The expression of TRPV4 in sensory neurons and its activation by changes in cell volume suggest that it detects osmotic and mechanical stimuli. TRPV4−/− mice show abnormal osmotic regulation and decreased acid and pressure nociception (not impaired touch or heat sensation).107,108 Although injection of hypotonic saline into rat hindpaws does not induce nociception, hypertonic saline causes pain, and pretreatment with an inﬂammatory mediator, PGE2, sensitizes responses to both stimuli.101,109 These effects require TRPV4, because they are prevented by TRPV4 deletion or downregulation. In a similar manner to PGE2, PAR2 agonists also increase the magnitude of the Ca2+ mobilization in response to hypotonic saline and 4αPDD in both human bronchial epithelial cells, which constitutively express both TRPV4 and PAR2, and in HEK cells heterologously expressing TRPV4 (Bunnett, unpublished). In slices of rat spinal cord, exposure to 4αPDD or hypotonic saline stimulates the release of SP and CGRP, which is potentiated by preexposure to PAR2 agonist. Hypotonic saline injected into mice paws does not produce any nociceptive behavior, but the combination of PAR2–AP and hypotonic saline increases the sensitivity to mechanical pain to a greater degree than PAR2–AP alone, indicating a sensitization of TRPV4 by PAR2 activation. TRPV4−/− mice do not show any increased sensitivity with either injection, indicating that both the mechanical hyperalgesia induced by PAR2 activation and the pain induced by hypotonic saline are mediated by TRPV4 (Bunnett, unpublished). Together, these ﬁndings suggest that PAR2 agonists can sensitize TRPV4 to induce hyperalgesia to mechanical stimuli, possibly by a mechanism that resembles those that mediate PAR2induced sensitization of TRPV1.

CONCLUSIONS AND FUTURE PERSPECTIVES PARs are widely expressed in the central and peripheral nervous systems, and a variety of protease sources can signal to neurons by cleaving PARs to regulate fundamentally important processes, which include pain transmission and neurogenic inﬂammation as well as neuronal survival, degeneration, neurite outgrowth, and release of neurotransmitters. Because PAR-activating proteases are generated or released during injury and inﬂammation, these mechanisms may be of considerable interest in treating diseases, and protease inhibitors and PAR antagonists could be useful therapeutic agents. However, there are major gaps in our understanding of the role of proteases and PARs in the nervous system. First, although several proteases have been characterized with the potential to activate neuronal PARs, the endogenous proteases that activate PARs in neuronal tissues under physiological and pathophysiological conditions have not been unequivocally identiﬁed. Second, the cellular mechanisms

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that transduce and regulate PAR signaling in neurons have not been deﬁned. Most information about PAR signaling and signal arrest derives from studies of cell lines, often transfected to express the receptor of interest, and it remains to be determined whether similar processes occur in neurons. Third, the mechanisms by which PARs regulate excitability of sensory nerves are not fully understood. Although it is well established that PAR2 agonists can sensitize TRPV1 and thereby induce thermal hyperalgesia, the signaling pathways responsible for this sensitization are not completely deﬁned. Moreover, the mechanisms of PAR2-induced mechanical hyperalgesia are not well characterized, and the mechanisms of the analgesic actions of PAR1 agonists are unknown. Fourth, although the roles of PAR1 and PAR2 have been studied in neurons, very little is known about the functions of PAR3 and PAR4, which are also present in the nervous system. Finally, the roles of proteases and PARs in experimental models of pain and neurogenic inﬂammation have not been fully explored, in part due to the lack of speciﬁc protease inhibitors and PAR antagonists. The use of such tools and of genetically modiﬁed animals will allow investigation of the contributions of proteases and PARs in experimental models of human diseases.

ACKNOWLEDGMENTS Research in the authors’ laboratory in this area is supported by NIH grants DK43207, DK57489, and DK39957, and by the Wellcome Trust.

REFERENCES 1. Southan, C., A genomic perspective on human proteases, FEBS Lett, 498, 214, 2001. 2. Macfarlane, S.R. et al., Proteinase-activated receptors, Pharmacol Rev, 53, 245, 2001. 3. Ossovskaya, V.S. and Bunnett, N.W., Protease-activated receptors: contribution to physiology and disease, Physiol Rev, 84, 579, 2004. 4. Vergnolle, N. et al., Proteinase-activated receptors: novel signals for peripheral nerves, Trends Neurosci, 26, 496, 2003. 5. Saito, T. and Bunnett, N.W., Protease-activated receptors: regulation of neuronal function, Neuromolecular Med, 7, 79, 2005. 6. Amadesi, S. et al., Protease-activated receptor 2 sensitizes the capsaicin receptor transient receptor potential vanilloid receptor 1 to induce hyperalgesia, J Neurosci, 24, 4300, 2004. 7. Dai, Y. et al., Proteinase-activated receptor 2–mediated potentiation of transient receptor potential vanilloid subfamily 1 activity reveals a mechanism for proteinaseinduced inﬂammatory pain, J Neurosci, 24, 4293, 2004. 8. Rasmussen, U.B. et al., cDNA cloning and expression of a hamster alpha-thrombin receptor coupled to Ca2+ mobilization, FEBS Lett, 288, 123, 1991. 9. Vu, T.K. et al., Molecular cloning of a functional thrombin receptor reveals a novel proteolytic mechanism of receptor activation, Cell, 64, 1057, 1991. 10. Nystedt, S. et al., Molecular cloning of a potential proteinase activated receptor, Proc Natl Acad Sci USA, 91, 9208, 1994.

Protease-Activated Receptors

435

11. Connolly, A.J. et al., Role of the thrombin receptor in development and evidence for a second receptor, Nature, 381, 516, 1996. 12. Xu, W.F. et al., Cloning and characterization of human protease-activated receptor 4, Proc Natl Acad Sci USA, 95, 6642, 1998. 13. Kahn, M.L. et al., A dual thrombin receptor system for platelet activation, Nature, 394, 690, 1998. 14. Andrade-Gordon, P. et al., Design, synthesis, and biological characterization of a peptide-mimetic antagonist for a tethered-ligand receptor, Proc Natl Acad Sci USA, 96, 12257, 1999. 15. Ferrell, W.R. et al., Essential role for proteinase-activated receptor-2 in arthritis, J Clin Invest, 111, 35, 2003. 16. Vu, T.K. et al., Domains specifying thrombin-receptor interaction, Nature, 353, 674, 1991. 17. Ishihara, H. et al., Protease-activated receptor 3 is a second thrombin receptor in humans, Nature, 386, 502, 1997. 18. Camerer, E. et al., Tissue factor– and factor X–dependent activation of proteaseactivated receptor 2 by factor VIIa, Proc Natl Acad Sci USA, 97, 5255, 2000. 19. Riewald, M. and Ruf, W., Mechanistic coupling of protease signaling and initiation of coagulation by tissue factor, Proc Natl Acad Sci USA, 98, 7742, 2001. 20. Riewald, M. et al., Activation of endothelial cell protease activated receptor 1 by the protein C pathway, Science, 296, 1880, 2002. 21. Nakanishi-Matsui, M. et al., PAR3 is a cofactor for PAR4 activation by thrombin, Nature, 404, 609, 2000. 22. O’Brien, P.J. et al., Thrombin responses in human endothelial cells. Contributions from receptors other than PAR1 include the transactivation of PAR2 by thrombincleaved PAR1, J Biol Chem, 275, 13502, 2000. 23. Emi, M. et al., Cloning, characterization, and nucleotide sequences of two cDNAs encoding human pancreatic trypsinogens, Gene, 41, 305, 1986. 24. Nyaruhucha, C.N. et al., Identiﬁcation and expression of the cDNA-encoding human mesotrypsin(ogen), an isoform of trypsin with inhibitor resistance, J Biol Chem, 272, 10573, 1997. 25. Wiegand, U. et al., Cloning of the cDNA encoding human brain trypsinogen and characterization of its product, Gene, 136, 167, 1993. 26. Kong, W. et al., Luminal trypsin may regulate enterocytes through proteinaseactivated receptor 2, Proc Natl Acad Sci USA, 94, 8884, 1997. 27. Hoogerwerf, W.A. et al., The proteinase-activated receptor 2 is involved in nociception, J Neurosci, 21, 9036, 2001. 28. Hoogerwerf, W.A. et al., Trypsin mediates nociception via the proteinase-activated receptor 2: a potentially novel role in pancreatic pain, Gastroenterology, 127, 883, 2004. 29. Cottrell, G.S. et al., Trypsin IV, a novel agonist of protease-activated receptors 2 and 4, J Biol Chem, 279, 13532, 2004. 30. Szmola, R. et al., Human mesotrypsin is a unique digestive protease specialized for the degradation of trypsin inhibitors, J Biol Chem, 278, 48580, 2003. 31. Sahin-Toth, M., Human mesotrypsin deﬁes natural trypsin inhibitors: from passive resistance to active destruction, Protein Pept Lett, 12, 457, 2005. 32. Steinhoff, M. et al., Agonists of proteinase-activated receptor 2 induce inﬂammation by a neurogenic mechanism, Nat Med, 6, 151, 2000. 33. Caughey, G.H., Mast cell chymases and tryptases: phylogeny, family relations, and biogenesis., in Mast cell proteases in immunology and biology, ed. Caughey, G. H. Marcel Dekker, New York, 1995, p. 305.

436

TRP Ion Channel Function in Sensory Transduction

34. Fiorucci, L. and Ascoli, F., Mast cell tryptase, a still enigmatic enzyme, Cell Mol Life Sci, 61, 1278, 2004. 35. Stead, R.H. et al., Intestinal mucosal mast cells in normal and nematode-infected rat intestines are in intimate contact with peptidergic nerves, Proc Natl Acad Sci USA, 84, 2975, 1987. 36. Compton, S.J. et al., Glycosylation and the activation of proteinase-activated receptor 2 (PAR2) by human mast cell tryptase, Br J Pharmacol, 134, 705, 2001. 37. Compton, S.J. et al., Glycosylation of human proteinase-activated receptor-2 (hPAR2): role in cell surface expression and signalling, Biochem J, 368, 495, 2002. 38. Mirza, H. et al., Mitogenic responses mediated through the proteinase-activated receptor-2 are induced by expressed forms of mast cell alpha- or beta-tryptases, Blood, 90, 3914, 1997. 39. Huang, C. et al., Evaluation of the substrate speciﬁcity of human mast cell tryptase beta I and demonstration of its importance in bacterial infections of the lung, J Biol Chem, 276, 26276, 2001. 40. Ishii, K. et al., Kinetics of thrombin receptor cleavage on intact cells. Relation to signaling, J Biol Chem, 268, 9780, 1993. 41. Hung, D.T. et al., The cloned platelet thrombin receptor couples to at least two distinct effectors to stimulate phosphoinositide hydrolysis and inhibit adenylyl cyclase, J Biol Chem, 267, 20831, 1992. 42. Klages, B. et al., Activation of G12/G13 results in shape change and Rho/Rhokinase–mediated myosin light chain phosphorylation in mouse platelets, J Cell Biol, 144, 745, 1999. 43. Offermanns, S. et al., Defective platelet activation in G alpha(q)-deﬁcient mice, Nature, 389, 183, 1997. 44. Bohm, S.K. et al., Molecular cloning, expression, and potential functions of the human proteinase-activated receptor-2, Biochem J, 314, 1009, 1996. 45. Corvera, C.U. et al., Mast cell tryptase regulates rat colonic myocytes through proteinase-activated receptor 2, J Clin Invest, 100, 1383, 1997. 46. Corvera, C.U. et al., Thrombin and mast cell tryptase regulate guinea-pig myenteric neurons through proteinase-activated receptors-1 and -2, J Physiol, 517, 741, 1999. 47. DeFea, K.A. et al., beta-arrestin-dependent endocytosis of proteinase-activated receptor 2 is required for intracellular targeting of activated ERK1/2, J Cell Biol, 148, 1267, 2000. 48. Ishii, K. et al., Inhibition of thrombin receptor signaling by a G-protein-coupled receptor kinase. Functional speciﬁcity among G-protein-coupled receptor kinases, J Biol Chem, 269, 1125, 1994. 49. Iaccarino, G. et al., Myocardial overexpression of GRK3 in transgenic mice: evidence for in vivo selectivity of GRKs, Am J Physiol, 275, H1298, 1998. 50. Tiruppathi, C. et al., G-protein-coupled receptor kinase-5 regulates thrombin-activated signaling in endothelial cells, Proc Natl Acad Sci USA, 97, 7440, 2000. 51. Paing, M.M. et al., beta-arrestins regulate protease-activated receptor-1 desensitization but not internalization or down-regulation, J Biol Chem, 277, 1292, 2002. 52. Hoxie, J.A. et al., Internalization and recycling of activated thrombin receptors, J Biol Chem, 268, 13756, 1993. 53. Bohm, S.K. et al., Mechanisms of desensitization and resensitization of proteinaseactivated receptor-2, J Biol Chem, 271, 22003, 1996. 54. Dery, O. et al., Trafﬁcking of proteinase-activated receptor-2 and beta-arrestin-1 tagged with green ﬂuorescent protein. Beta-arrestin-dependent endocytosis of a proteinase receptor, J Biol Chem, 274, 18524, 1999.

Protease-Activated Receptors

437

55. Trejo, J. et al., Protease-activated receptor-1 down-regulation: a mutant HeLa cell line suggests novel requirements for PAR1 phosphorylation and recruitment to clathrin-coated pits, J Biol Chem, 275, 31255, 2000. 56. Roosterman, D. et al., Rab5a and rab11a mediate agonist-induced trafﬁcking of protease-activated receptor 2, Am J Physiol Cell Physiol, 284, C1319, 2003. 57. Wang, Y. et al., Down-regulation of protease-activated receptor-1 is regulated by sorting nexin 1, Mol Biol Cell, 13, 1965, 2002. 58. Jacob, C. et al., c-Cbl mediates ubiquitination, degradation, and down-regulation of human protease-activated receptor 2, J Biol Chem, 280, 16076, 2005. 59. Weinstein, J.R. et al., Cellular localization of thrombin receptor mRNA in rat brain: expression by mesencephalic dopaminergic neurons and codistribution with prothrombin mRNA, J Neurosci, 15, 2906, 1995. 60. Junge, C.E. et al., Protease-activated receptor-1 in human brain: localization and functional expression in astrocytes, Exp Neurol, 188, 94, 2004. 61. de Garavilla, L. et al., Agonists of proteinase-activated receptor 1 induce plasma extravasation by a neurogenic mechanism, Br J Pharmacol, 133, 975, 2001. 62. Smith-Swintosky, V.L. et al., Protease-activated receptor-2 (PAR-2) is present in the rat hippocampus and is associated with neurodegeneration, J Neurochem, 69, 1890, 1997. 63. D’Andrea, M.R. et al., Characterization of protease-activated receptor-2 immunoreactivity in normal human tissues, J Histochem Cytochem, 46, 157, 1998. 64. Wang, H. et al., Four subtypes of protease-activated receptors, co-expressed in rat astrocytes, evoke different physiological signaling, Glia, 37, 53, 2002. 65. Kaufmann, R. et al., The two-receptor system PAR-1/PAR-4 mediates alpha-thrombin-induced [Ca(2+)](i) mobilization in human astrocytoma cells, J Cancer Res Clin Oncol, 126, 91, 2000. 66. Ubl, J.J. et al., Co-existence of two types of [Ca2+]i-inducing protease-activated receptors (PAR-1 and PAR-2) in rat astrocytes and C6 glioma cells, Neuroscience, 86, 597, 1998. 67. Linden, D.R. et al., Agonists of proteinase-activated receptor 2 excite guinea pig ileal myenteric neurons, Eur J Pharmacol, 431, 311, 2001. 68. Gao, C. et al., Serine proteases excite myenteric neurons through protease-activated receptors in guinea pig small intestine, Gastroenterology, 123, 1554, 2002. 69. Reed, D.E. et al., Mast cell tryptase and proteinase-activated receptor 2 induce hyperexcitability of guinea-pig submucosal neurons, J Physiol, 547, 531, 2003. 70. Green, B.T. et al., Intestinal type 2 proteinase-activated receptors: expression in opoidsensitive secretomotor neural circuits that mediate epithelial ion transport, J Pharmacol Exp Ther, 295, 410, 2000. 71. Cuffe, J.E. et al., Basolateral PAR-2 receptors mediate KCl secretion and inhibition of Na+ absorption in the mouse distal colon, J Physiol, 539, 209, 2002. 72. Vergnolle, N. et al., Proteinase-activated receptor-2 and hyperalgesia: a novel pain pathway, Nat Med, 7, 821, 2001. 73. Barbara, G. et al., Activated mast cells in proximity to colonic nerves correlate with abdominal pain in irritable bowel syndrome, Gastroenterology, 126, 693, 2004. 74. Minn, A. et al., Enhanced GFAP expression in astrocytes of transgenic mice expressing the human brain-speciﬁc trypsinogen IV, Glia, 22, 338, 1998. 75. Chraibi, A. et al., Protease modulation of the activity of the epithelial sodium channel expressed in Xenopus oocytes, J Gen Physiol, 111, 127, 1998. 76. Krishtal, O., The ASICs: signaling molecules? modulators? Trends Neurosci, 26, 477, 2003.

438

TRP Ion Channel Function in Sensory Transduction

77. Sutherland, S.P. et al., Acid-sensing ion channel 3 matches the acid-gated current in cardiac ischemia-sensing neurons, Proc Natl Acad Sci USA, 98, 711, 2001. 78. Poirot, O. et al., Selective regulation of acid-sensing ion channel 1 by serine proteases, J Biol Chem, 279, 38448, 2004. 79. Nicole, O. et al., The proteolytic activity of tissue-plasminogen activator enhances NMDA receptor-mediated signaling, Nat Med, 7, 59, 2001. 80. Asfaha, S. et al., Proteinase-activated receptor-1 agonists attenuate nociception in response to noxious stimuli, Br J Pharmacol, 135, 1101, 2002. 81. Cesare, P. et al., Ion channels gated by heat, Proc Natl Acad Sci USA, 96, 7658, 1999. 82. Numazaki, M. et al., Direct phosphorylation of capsaicin receptor VR1 by protein kinase C epsilon and identiﬁcation of two target serine residues, J Biol Chem, 277, 13375, 2002. 83. Vellani, V. et al., Protein kinase C activation potentiates gating of the vanilloid receptor VR1 by capsaicin, protons, heat, and anandamide, J Physiol, 534, 813, 2001. 84. Mandadi, S. et al., Activation of protein kinase C reverses capsaicin-induced calciumdependent desensitization of TRPV1 ion channels, Cell Calcium, 35, 471, 2004. 85. Prescott, E.D. and Julius, D., A modular PIP2 binding site as a determinant of capsaicin receptor sensitivity, Science, 300, 1284, 2003. 86. Chuang, H.H. et al., Bradykinin and nerve growth factor release the capsaicin receptor from PtdIns(4,5)P2-mediated inhibition, Nature, 411, 957, 2001. 87. Zhang, X. et al., NGF rapidly increases membrane expression of TRPV1 heat-gated ion channels, Embo J, 24, 4211, 2005. 88. Kawao, N. et al., Capsazepine inhibits thermal hyperalgesia but not nociception triggered by protease-activated receptor-2 in rats, Jpn J Pharmacol, 89, 184, 2002. 89. Kawabata, A. et al., Capsazepine partially inhibits neurally mediated gastric mucus secretion following activation of protease-activated receptor 2, Clin Exp Pharmacol Physiol, 29, 360, 2002. 90. Khasar, S.G. et al., A novel nociceptor signaling pathway revealed in protein kinase C epsilon mutant mice, Neuron, 24, 253, 1999. 91. Rozengurt, E. et al., Protein kinase D signaling, J Biol Chem, 280, 13205, 2005. 92. Tan, M. et al., Thrombin rapidly induces protein kinase D phosphorylation, and protein kinase C delta mediates the activation, J Biol Chem, 278, 2824, 2003. 93. Wang, Y. et al., Interaction between protein kinase Cmu and the vanilloid receptor type 1, J Biol Chem, 279, 53674, 2004. 94. Malmberg, A.B. et al., Diminished inﬂammation and nociceptive pain with preservation of neuropathic pain in mice with a targeted mutation of the type I regulatory subunit of cAMP-dependent protein kinase, J Neurosci, 17, 7462, 1997. 95. Bhave, G. et al., cAMP-dependent protein kinase regulates desensitization of the capsaicin receptor (VR1) by direct phosphorylation, Neuron, 35, 721, 2002. 96. Mohapatra, D.P. and Nau, C., Desensitization of capsaicin-activated currents in the vanilloid receptor TRPV1 is decreased by the cyclic AMP-dependent protein kinase pathway, J Biol Chem, 278, 50080, 2003. 97. Lopshire, J.C. and Nicol, G.D., The cAMP transduction cascade mediates the prostaglandin E2 enhancement of the capsaicin-elicited current in rat sensory neurons: whole-cell and single-channel studies, J Neurosci, 18, 6081, 1998. 98. Rathee, P.K. et al., PKA/AKAP/VR-1 module: a common link of Gs-mediated signaling to thermal hyperalgesia, J Neurosci, 22, 4740, 2002. 99. Coelho, A.M. et al., Proteinases and proteinase-activated receptor 2: a possible role to promote visceral hyperalgesia in rats, Gastroenterology, 122, 1035, 2002.

Protease-Activated Receptors

439

100. Liedtke, W. et al., Vanilloid receptor-related osmotically activated channel (VR-OAC), a candidate vertebrate osmoreceptor, Cell, 103, 525, 2000. 101. Alessandri-Haber, N. et al., Hypotonicity induces TRPV4-mediated nociception in rat, Neuron, 39, 497, 2003. 102. Guler, A.D. et al., Heat-evoked activation of the ion channel, TRPV4, J Neurosci, 22, 6408, 2002. 103. Clapham, D.E. et al., International Union of Pharmacology. XLIII. Compendium of voltage-gated ion channels: transient receptor potential channels, Pharmacol Rev, 55, 591, 2003. 104. Watanabe, H. et al., Activation of TRPV4 channels (hVRL-2/mTRP12) by phorbol derivatives, J Biol Chem, 277, 13569, 2002. 105. Watanabe, H. et al., Anandamide and arachidonic acid use epoxyeicosatrienoic acids to activate TRPV4 channels, Nature, 424, 434, 2003. 106. Pedersen, S. et al., Hypotonic cell swelling induces translocation of the alpha isoform of cytosolic phospholipase A2 but not the gamma isoform in Ehrlich ascites tumor cells, Eur J Biochem, 267, 5531, 2000. 107. Liedtke, W. and Friedman, J.M., Abnormal osmotic regulation in trpv4−/− mice, Proc Natl Acad Sci USA, 100, 13698, 2003. 108. Suzuki, M. et al., Impaired pressure sensation in mice lacking TRPV4, J Biol Chem, 278, 22664, 2003. 109. Alessandri-Haber, N. et al., TRPV4 mediates pain-related behavior induced by mild hypertonic stimuli in the presence of inﬂammatory mediator, Pain, 2005.

Index A Accessory olfactory bulb (AOB), 46, 47 Acid-evoked pain, 153 Acids, see Protons/hydrogen ions/acids/pH Acid-sensing ion channels, 73–74, 430 Acoustic stimuli, 147; see also Auditory signals/hearing Acrolein, 155, 156, 170 Acrosome, 48, 196 Actin polycystins and, 259 stereocilia, 228 TRP-PLC signaling pathway, 15 Aδ ﬁbers, 180, 276 Adenine nucleotides, 205; see also ATP Adenosine triphosphate (ATP), see ATP Adenylyl cyclase, 191 ADPKD, 196 ADP ribose, 4, 7, 355 AG-3-5, see Icilin (AG-3-5) Aggression, pheromone sensing, 46, 47, 50 Airway, see Respiratory tract AKAP450, 231 AKT, 90–91 Aldehydes, TRPA1 sensitivity, 154, 170 Algae, 232, 260 Allicin, 154, 156, 170, 279 Allodynia, cold, 158 Allosteric model temperature sensitivity, 297–299 TRPM8, 185 Allyl isothiocyanate, 170, 280; see also Mustard oils Alpha kinase, TRPM7, 355 Alpha toxin, 191, 260–261 AMG 9810 analogue, 93 Amiloride, 166, 190, 365 Amino acid alignment, 7 Amino acid detection, pheromone sensing, 49 Amino acid homology, TRPC family, 364 Amino acid identity, TRPM5, 204 Amino terminus TRPC2, 48, 49 TRPC protein–protein interactions, 334, 336, 337–338

TRPV1, 77 TRPV4, 114, 115, 142, 143 AMPA, 322 AMPA-type receptors, 18 AMPPNP, 351 Amygdala, 46 Anandamide, 7, 131 TRPA1, 155–156 TRPV1, 70, 71, 73, 97 TRPV4, 119, 144, 398 Ankyrin domain, TRPV4, 130 Ankyrin repeats, 351 genomic structures, 8, 10, 13 OCR-2, 250 oligomerization of cytosolic domains, 356 receptor structure, 321 structure, 353–355 TRPA1, 3, 163, 165, 235 TRPC2, 48 TRPC family, 334, 335 TRPP2, 198 TRPV1, 70 TRPV2, 390 TRPV4, 142, 143 Ankyrin subfamily (TRPA), see TRPA (subfamily) ANKTM1, see TRPA1 (ANKTM1) Annexin 2, 327 Antennal chordotonal organs, 229–230 Antidiuretic hormone, 117, 126, 309 Antinociceptive effects, TRPV1, 89, 90 Antisense technology neuron pathﬁnding/migration, 58 TRPA1, 158 TRPC1, 369 TRPC family, 332 TRPV2, 368 AP-1, 192, 371 Apes pheromone sensing, molecular biology of, 50 TRPC2 as pseudogene in, 50 vomeronasal system, 47 Apoptosis, TRPV1 and, 96–97 Aquaporin-2 (AQP-2), 322 Aquaporin-5 (AQP-5), 308 Arachidonic acid

441

442

TRP Ion Channel Function in Sensory Transduction

EETs, see Epoxyeicosatrienoic acid (EET) metabolites of arachidonic acid protease-activated receptors, 426, 433 TRPV1, 70, 71, 77 TRPV4, 114, 144, 145, 307, 398 ciliated epithelia, 417 endothelial cells, 382–383 and ﬂow-induced vasodilatation, 384 gating, molecular mechanisms, 117 molecular mechanisms of gating, 119 vascular endothelium, 371, 378, 382–383, 384 N-Arachidonoyldopamine (NADA), 76, 88 N-Arachidonoyl-phosphatidylethanolamine (NAPE), 70, 73 Arginine vasopressin, 117, 126, 309 Arista, 229 Arrestins, 426, 427 Arrhenius activation energy, 289 N-Arylcinnamides, 93 ASH sensory neurons, 310, 311–312, 313 atonal mutants, 216, 228 ATP, 7, 158, 260–261 protease-activated receptor sensitization of TRPV channels, 430, 431 TRPM4, endothelial, 380 TRPM4/TRPM5 sensitivity, 205 TRPV1 modulation, 76 vascular endothelium, 370 ATP-2, 258, 260 ATP synthase, 260, 261 Auditory signals/hearing, 222 ciliary localization, 259 Drosophila, see Drosophila, hearing and proprioception mechanosensitivity and, 303 TRPA1, 164–169, 171, 223 channel properties of TRPA1 versus hair cell transducer, 166–167 inner ear, distribution in, 164–165 inner ear, functional tests in, 165–166 sensory ganglia, 169 TRPV4 and, 4, 144, 147 Type I sensory neurons, 217 Autonomic nervous system, TRPA1, 164 Autosomal polycystic kidney disease, see Polycystic kidney disease Axon pathﬁnding, TRP channels and, 4, 55–63 calcium signaling in growth cone turning, 56–57 function in nerve guidance, 57–61 gating mechanisms by guidance factors, 59 growth regulation, TRPC5 and, 57–58 mechanism of contributions to guidance signaling, 59–61 pathﬁnding process, TRPC and, 58–59

overview of nerve pathﬁnding, 55–56 possible roles of channels in neuronal migration, 61–63 calcium signal in, 61 mechanosensation in migrating neurons, 61–63 thermosensation for migration, 63

B Bacterial KcsA potassium channel, 114, 115, 119, 120 Barium, 133, 190 Basic ﬁbroblast growth factor (bFGF), 370 BCTC, 93 TRPM8 antagonists, 183 TRPV1 modulation, pharmacological overlap, 96 Beethoven, 233 Behavioral mediators, TRPC2 and pheromone detection in mammals, 45–51; see also TRPC2, and pheromone detection in mammals Benzyl isothiocyanate, 153, 170 Biophysics temperature sensitivity, see Thermosensation, channel gating biophysics TRPV4 properties, 136 BKCa channels, 366 Bladder, 73, 91, 94 Blood pressure, 377; see also Vasodilation Blood vessels anandamide triphasic response, 155 ROCE and SOCE mechanisms, 23 TRPA1, cannabinoids and, 156 TRPM4 function, 208 TRPP2, 194 TRPV1 localization, 94 TRPV2, 107, 389–394 endothelium, 393 smooth muscle, 390–394 TRPV4, 113, 143 vascular function, endothelial cells, 369–372 calcium entry, 369–371 channel presence in endothelium, 369 oxidative stress sensors, 372 proliferative response to hypoxia, 371 regulation of capillary permeability, 371 temperature change, response to, 371 TRPV4 role in mechanotransduction of shear stress, 377–386 vascular function, smooth muscle cells, 363–369 pressure-induced channel activation, 366–368

Index store-operated channel activation, 368–369 vasoconstriction agonist-induced channel activation, 363–366 vasodilator response to EETs mediated by channel activation, 366 B lymphocytes, 23, 34 Botulinum neurotoxins, 325, 327, 328 Bradykinin, 19, 75, 76, 158, 183 protease-activated receptor sensitization of TRPV channels, 430, 431 targets of, 96 TrkA receptor, 158 TRPA1, 170, 171 TRPV1, 77, 90, 274 and vascular endothelium, 370 Brain, see also Central nervous system neuron pathﬁnding/migration, 55–63 TRPC2, 48 TRPM3, 146 TRPV4, 126 Brain-derived neurotrophic factor (BDNF), 4, 56, 58, 59, 60, 61, 326 Breast cancer, TRPM8, 185 Breast pain, 97

C CABP1, 342 Cadmium, 365 Caenorhabditis elegans, 232, 234, 235, 380 OCR-2 and OSM-9 function of, 246–247 speciﬁcity of signaling pathways in different sensory neurons, 247–251 OSM-9, 116, 126, 127, 129, 234, 260, 276, 380, 397, 398, 432 mechanotransduction, 222 osmo- and mechanotransduction, channel functions in, 303–304, 310–313 TRPA1, 171 TRPA1 orthologue, 164 TRP channels, 198 TRPP2 and PKD1 heteromeric complexes, 196 TRPV channels in serotonergic neurons, 243–252 conservation of serotonergic system, 244–246 speciﬁcity of signaling pathways in different sensory neurons, 247–251 vomeronasal neuron signal transduction similarity, 46 Caenorhabditis elegans, mating and fertilization, 257–264

443 TRPC channels, 262–264 questions and future directions, 264 TRP-3 and sperm-egg interactions, 262–263 TRP-3 translocation during sperm activation, 263–264, 325 TRPP, 257–261 future directions, 261 intraﬂagellar transport of TRPV but not TRPP in cilia, 260 kinesin KLP-6 regulation, 259–260 PLAT binding partners, 260–261 polycystin ciliary localization and ADPKD, 257–259 Calcineurin, 258, 261 FK506 binding and, 15 TRPC, 14 TRPV1, 76–77, 89 dissimilar structure-activity relationships, 92 intracellular compartment translocation, 91 Calcitonin gene-related peptide (CGRP), 428, 429, 431, 433 Calcium, 2, 7, 234, 280, 323–324 blood vessels, see Blood vessels and cell cycle progression, 105–106 neuron pathﬁnding/migration gating mechanisms by guidance factors, 59 gradient, 56–57 mechanisms, 59, 60–61 possible roles, 61 regulation of nerve growth, 57–58 temperature sensitivity, 63 TRPC, 58 odorant, osmotic, and mechanical stimuli, 312 ROCE SOCE conundrum, 2 selectivity/nonselectivity of channels, 4 TRPA1 cannabinoid receptors, 157 hair cell transducer, 167, 168 TRPC family activation mechanisms, 2–3, 37 channel translocation to plasma membrane and, 17–18 GPCR-Gq activation to voltage-gated calcium channel activation, 18 signal transduction, 14 tyrosine kinases in voltage-gated calcium channel-independent calcium inﬂux, 19 TRPM4/TRPM5 sensitivity, 205–206, 208–209 TRPM8, 178, 183–184 TRPP2 and TRPP2-related channels, 190, 196–197, 198–199 TRPV1, 87 CAMKII and, 76–77 dissimilar structure-activity relationships, 91–92 signal transduction, 70, 71

444

TRP Ion Channel Function in Sensory Transduction

TRPV2, 106–107, 111, 112 TRPV4, 127, 133–134 activation and pharmacological properties, 143–144 calmodulin binding sites, 115, 116 channel properties, 135–136 mechanosensitivity, 129, 131 osmotic changes and, 131, 142 TRPV4 gating, molecular mechanisms channel selectivity, 119 mechanosensory, 117 voltage dependence, see Voltage-dependent calcium channels Calcium-binding protein 1 (CABP1), 342 Calcium release activated calcium current (Icrac), 3, 17, 35 Calmodulin/calmodulin binding FK506 binding and, 15 genomic structures, 8, 12 receptor structure, 321 TRPC2 interaction, 49 TRP channels, 2 TRPC protein–protein interactions, 336, 340, 341–342 TRPV1 modulation, 75, 76–78 TRPV1 structure, 70 TRPV4 structure, 115, 116, 130, 133, 134, 142, 144 Calmodulin-dependent protein kinase II (CAMKII), 76–77, 90–91 Calyculin A, 15 CAMKII (calmodulin-dependent protein kinase II), 76–77, 90–91 CAM kinase, 432 Camphor, 280 Cancer cells TRPM8, 185 TRPV1, 96–97 TRPV2, IGF regulation, 106 Cannabinoids, 280 TRPA1 activation, 155–157 TRPV1 activation, 70, 71, 88 Canonical subfamily (TRPC), see TRPC (subfamily) Capacitative calcium entry (CCE), 3; see also Store-operated calcium entry (SOCE) Capillary permeability, regulation of, 371 Capsaicin, 7, 234, 280, 310 binding sites, 352 and cannabinoid-induced hypotension, 156 diversity of behavior, structure-activity relationships, 86–87 protease-activated receptor agonist synergy, 432 TRPA1 probes, 152–155 TRPV1 activation, 70, 72–73, 88, 217

NGF and inﬂammatory hyperalgesia, 90 pathological conditions, changes of expression under, 97 pH and, 74 phosphorylation and, 76–77, 91 serum steroids and, 95 sulfhydryl group oxidation state and, 78 Capsaicin receptor cDNA, 3 Capsazepine, 89, 93 TRPA1, 156 TRPM8, 183, 185 Carbachol, 20 Carboxy terminus oligomerization of, 356 TRPC2, 48 TRPC protein–protein interactions, 336, 340, 342 TRPM3, 145 TRPP2, 193 TRPP2-like proteins, 190 TRPV1, 70, 77 TRPV4, 115, 116, 133, 142 Cardiovascular system, 377; see also Blood vessels; Heart anandamide and, 155 TRPV1 in cardiomyocytes, 94 TRPV2, 389–394 expression, 391 mechanisms of mechanoactivation, 394 mechanosensation, 391–394 Carotid artery, TRPV4, 380–381 Casein kinase 2, 194, 260, 261 CaT, 3 CaT1, see TRPV6 (CaT1, EcAC2) CaT2, see TRPV5 (CaT2, ECaC1) Cathepsin G, 426 Caveolin-1, 336, 337 c-Cbl, 428 CCR1 (chemokine receptor), TRPV1 modulation, 90 Cell compartments, see Trafﬁcking/translocation/sorting Cell culture neuron pathﬁnding/migration, 58, 59, 62 osmo- and mechanotransduction, channel functions in, 306–310 TRPP2 and PKD1 coexpression, 195 tyrosine phosphorylation, role in TRPC function, 19–20 Cell cycle, TRPV2, calcium entry requirement, 105–106 Cellular expression, see Tissue expression/localization/distribution Cellular function, TRPV4 and, 144–145 Cell volume

Index and cytoskeleton, 141 regulation of, see Volume regulation Central nervous system, see also Dorsal root ganglia (DRG) cerebral blood ﬂow, 208, 367, 380–381 neuron pathﬁnding/migration, see Axon pathﬁnding, TRP channels and ROCE and SOCE mechanisms, 23 TRPC2 and pheromone detection in mammals, 46 TRPM3, 146 TRPV1, 73, 94 TRPV4, 113, 143 Ceramides, TRPM3 activation, 147 Cerebellum neuron pathﬁnding/migration, 4, 58, 59, 61, 62 TRPM3, 146 Cerebral blood ﬂow, 208, 367, 380–381 Cervical carcinoma, 97 Cesium current, 109, 110 C-ﬁbers, 152 ethanol and, 88 mechanicoheat nociceptors, 271 TRPA1, 169 TRPM8 expression, 180 TRPV1, 306 CGRP, 279 Channel-activating proteases, 430 Channel gating, see Gating Chemokines TRPV1 modulation, 90–91 Chemosensation Caenorhabditis elegans OCR-2 and OSM-9, 243–252 vomeronasal neuron signal transduction similarity, 46 neuron pathﬁnding/migration, 56, 58–59, 60 TRPA1, 151 Chlamydomonas, 232, 260 CHO cells TRPP2 and PKD1 coexpression, 195 TRPV2 expression, 306–307, 308, 393 Chordotonal neurons, 217, 234; see also Drosophila, hearing and proprioception Choroid plexus, 126, 146 Chromosomal loci, 4 Chromosomal repeats, 12–13 Chymotrypsin, 430 Cilia/ciliated epithelium, 196, 229 Caenorhabditis elegans polycystin localization, 257–259 TRPV intraﬂagellar transport, 257–259 Drosophila

445 hearing and proprioception, 227–228, 229, 230, 231 nan and iav, 234 unc mutants, 231 touch transduction, 236, 237 TRPA1, 165 TRPP2, 196–197 TRPV4, 413–418 Ciliary beat frequency, 416–417 Cinnamaldehyde, 154, 156, 170, 222, 280 CIRB (calmodulin and IP3R binding site), 340 Circadian clock, 4 Circulatory organs, see Cardiovascular system Circumventricular organs, TRPV4, 143 Ck protein, 233 CMR1, see TRPM8 (CMR1) Cobalt, 106 Cochlea hair cells, see Hair cells stereocilia, 228 Coding sequences, 4 Coexpression/colocalization KLP-6 with LOV-1/PC-1 and OKD-2/PC2, 259 PAR2 and β-arrestins, 427 PKD-2 and ATP synthase, 261 TrkA, 180 TrkA receptor, 169 TRPA1 and TrkA receptor, 169 and TRPV1, 154, 157, 279 TRPM8 and TrkA, 180 TRPP2 and PKD1, 195 TRPV1, 88, 96, 154, 157, 279 and v-SNARE protein, 327, 328 TRPV4 with MAP7, 116 Coiled coil domain, 352 polycystin-2, 258–259 protein–protein interactions, 336 TRPC2, 48 TRPC family, 334 TRPP2-like proteins, 190 Cold allodynia, 158 Cold receptors/sensitivity, 4, 7 temperature sensors in mammals, 217 thermoTRP channel gating biophysics, 289 TRPA1, 157–158, 222 TRPA1 and, 170–171 TRPM8, 4, 177–186 Colon, TRPM4, 204 Colon cancer, TRPM8, 185 Combinatorial encoding of temperature response, 218–220 Commissural axons, xTRPC1 and, 58–59 Competence factors, cell cycle, 105–106 Conformational changes, TRPV4 channel, 120

446

TRP Ion Channel Function in Sensory Transduction

Conformational coupling TRPC family activation mechanisms, 15–17 TRPC protein–protein interactions, 336, 341–342 CoolactP, 182 Cooling Agent 10, 182 COS-7 cells, 19, 20, 22, 24 Cough response TRPA1, 159 TRPV1 and, 97 CRAC channels, 323–324 Icrac, 3, 17, 35 TRPM4 function, 208, 209 Crinkled mutant, 233 c-src, 19 CTPC TRPM8 antagonists, 183 TRPV1 modulation, 96 Current activation/deactivation, temperature sensitivity, 295 Cyclic AMP, neuron pathﬁnding/migration, 60 Cyclic AMP-dependent protein kinase, see Protein kinase A (PKA) Cyclic GMP, neuron pathﬁnding/migration, 60 Cyclophosphamide, 155 Cyclopiazonic-induced SOC-mediated calcium entry, 325 Cyclosporin, 15 Cysteine residues, TRPV1, 70, 77–78 Cystic ﬁbrosis transmembrane conductance regulator (CFTR), 322 Cystitis, 91, 155 Cytochalasin D, 110 Cytochrome P450 epoxygenase, 433 endothelial function, 378 TRPV4, 117, 119, 131, 144, 307 Cytokine secretion, TRPM4 and, 208 Cytoskeleton cell volume and, 141 neuron pathﬁnding/migration, 62 polycystins and, 259 protein–protein interactions, 335 touch transduction, 235 TRPA1, 166 TRPA1 ankyrin repeats and, 163 TRPP2 and, 198 TRP-PLC signaling pathway, 15 TRPV2 and, 110, 394

D dDAVP, 309 Degenerin/epithelial sodium channel, 235 Dehydroepiandrosterone, TRPV1 modulation, 95 Dendritic cap, 231 Development

polycystins and, 260 TRPV1 localization, 94 Diacylglycerol (DAG) blood vessels endothelial function, 378 smooth muscle cells, 364 vasoactive substances and, 363 protease-activated receptors, 426 TRPC2 transduction, 49 TRPC family activation mechanisms, 17, 20, 21, 22, 31, 33 TRPC signal transduction, 14 TRPM8, 184 TRPV1, 77, 90 TRPV4 mechanosensitivity, 116–117 molecular mechanisms of gating, 118 Diallyl disulﬁde, 154, 155, 156 Dihydropyridine, 16 Direct chromosomal repeats, 12–13 Distortion product otoacoustic emissions (DPOAEs), 165 Dithiothreitol, 77–78 Dorsal root ganglia (DRG) neuron pathﬁnding/migration, 62 osmo- and mechanotransduction, channel functions in, 309, 310 protease-activated receptors, 431 protease signaling in nervous system, 428 temperature sensing, 217, 288, 289 TRPA1, 157, 163, 169 TRPM8, 178, 180, 279 TRPV1, 72, 73, 94, 274 TRPV2, 275, 276 TRPV4, 113 d-plp mutants, 231 Drosophila, 31, 33, 261, 364, 379 ciliary localization, 259 coiled coil domain, 334–335 dynamic translocation of photoreceptor cells, 325 osmo- and mechanotransduction, channel functions in, 303, 304, 313 painless gene, 171, 280–281 phototransduction, vomeronasal neuron signal transduction similarity, 46 protein–protein interactions, 335 binding partner effects on channel localization and function, 336 hetero and homomeric ion channels, 333 signaling partnerships, 342 TRPC signal transduction, 15 TRP families, 198, 322–323 mammalian homologues, 3 sequence similarity, 142 trpl discovery, 2

Index TRPN channels, 152 ankyrin repeats, 354 Drosophila, hearing and proprioception, 227–237 chordotonal organs, antennal and nonantennal, 229–233 gene products in mechanosensation, 231–233 gene products in mechanosensation, auditory and sensory transduction, 231–233 gene products in mechanosensation, auditory but not sensory transduction, 233 scolopidium structure, 230–231 TRPV channels, Nanchung and inactive, 234–237 Drosophila, nociception, 213–223 combinatorial encoding of temperature response, 218–220 innocuous versus noxious touch detection, distinct pathways for, 216–217 mammalian TRPA1 channel, multiple elicitors of burning pain, 222 mechanotransduction by TRP channels, 222–223 model development, 214 multiple heat activated TRPA channels in ﬂies, 218 temperature sensors, 272 temperature sensors in mammals, 217–218 thermosensory neurons, functions of highthreshold and low-threshold types, 221 TRPA channel mutations, increased mechanical and thermal nociception thresholds, 214–216 Drosophila, TRPC protein–protein interactions, 340 DT40, 34 Dysaesthesias, TRPV1, 306

E EB1 protein, Drosophila (DmEB1), 233 EBP-50, 335 ECaC, 3 ECaC1, see TRPV5 (CaT2, ECaC1) ECaC/CaT channels calcium selectivity, 4 discovery of, 3 genomic structures, 12, 13 TRPV5, see TRPV5 (CaT2, ECaC1) TRPV6, see TRPV6 (CaT1, ECaC2) EETs, see Epoxyeicosatrienoic acid (EET) metabolites of arachidonic acid Egg jelly receptor, 190, 191, 195, 196 Eicosanoids, TRPV1 modulation, 70, 71 Elastase, 426 Electrophysiological calcium entry, 3

447 TRPC family, 35 voltage dependence, see Voltage-dependent calcium channels Embryonic development polycystins and, 260 TRPP2 in, 197 Endocannabinoids, see Anandamide; Cannabinoids Endocytosis protease-activated receptors and, 426 TRPV2, 109, 110 Endogenous ligands, TRPV1 regulation, 87–89 Endoplasmic reticulum, 319 channel trafﬁcking, see Trafﬁcking/translocation/sorting SERCA, 2 TRPC2 splice variants, 49 TRPM8, 185 TRPP2, 193, 194 TRPV2, 107, 108, 111 Endothelin, 363 Endothelium TRPC trafﬁcking, 339 TRPV4, 113, 119, 126 osmolarity sensing, 117 vascular function, 369–372 calcium entry, 369–371 channel presence in endothelium, 369 oxidative stress sensors, 372 proliferative response to hypoxia, 371 regulation of capillary permeability, 371 temperature change, response to, 371 TRP channels, 362 Endothelium-derived hyperpolarizing factor (EDHF), 378, 385 Endovanilloids, TRPV1 ligands, 87–89 EP1 (prostaglandin receptor), TRPV1 modulation, 90 Ephrins, 63 Epidermal growth factor (EGF), 20, 22, 34, 326 Epidermal growth factor (EGF) receptor, 426 Epithelial calcium transporters, 3 Epithelial sodium channel, 322 Epithelium, channel-activating proteases, 430 Epitope mapping, 7 Epoxyeicosatrienoic acid (EET) metabolites of arachidonic acid, 117, 119, 130, 131, 144, 145, 280 blood vessels endothelium, 370, 371, 378 smooth muscle, 362, 365, 366 TRPV4 activation, 398 Epoxygenase metabolites, 114, 119 ERK (extracellular signal-regulated protein kinase), 90–91, 192, 426

448

TRP Ion Channel Function in Sensory Transduction

ERM family of proteins, 15 Ethanol, as vanilloid, 88–89 Ethanolamide, 70 Ethanolamines, see Anandamide Eucalyptol, 289 Eugenol, 154, 170 Euphorbia resinifera, 73, 87 Evodiamine, 86 Evolutionary conservation, TRPV4, 303–304 Excitation coupling model, 16 Exocytosis, 320, 321–322, 325, 326, 327 translocation of channels, 327, 328 TRPV2, 109 Exons, 9, 10, 11 GenBank data, 4, 5–6 genomic structures, 8 Expression sequence tags (ESTs) TRPC2, 48 TRPM4, 203 Extracellular factors, neurite extension, 57 Eyring’s transition state theory, 296–297 Ezrin-Radixin-Moesin (ERM) family of proteins, 15, 335

F Fat tissue, TRPV2, 107 Fatty acid amide hydrolase (FAAH), TRPV1 activation, 70 Fatty acid desaturase (fat-3), 398 Femoral chordotonal organ (FCO), 236–237 Fetal brain, TRPM3, 146 Fibroblasts, 19, 20, 22, 106–110, 112 Filopodia, 62 FK506, 15 FKBP12, 15 FKBP52, 336 Flagella, 232, 260 Flow dynamics, TRPV4 mechanosensitivity, 117 Fluids and electrolytes, TRPV4 and, 117 fos, 431 FrescolatMGA, 182 FrescolatML, 182 Fura-2, 2, 21 fyn-deﬁcient cells, 19, 20, 21, 22

G GABA, 305, 322, 428 Gadolinium, 22, 34, 62, 143, 166–167, 171, 365, 382, 384, 390, 417 Gα13, 208 Garlic derivatives, 222, 279, 288 TRPA1 activation, 154, 155, 156 Gastroesophageal reﬂux, 97

Gastrointestinal disease, TRPV1 in, 97 Gastrointestinal tract CaT discovery, 3 TRPM4/TRPM5, 204 TRPM8, 185 TRPV1, 94, 97, 305–306 TRPV2, 107 TRPV4, 308 Gating protein–protein interactions, 339–341 structure and, 353 thermoTRP channel gating biophysics, 287–299 TRPA1, 166 TRPV2, 109, 111 TRPV4, molecular mechanisms, 113–121 GenBank, 4, 5–6 Gene expression, TRPV1, 94, 95, 97 Genes and gene products, TRPC, 4–13 direct chromosomal repeats, 12–13 genomic structures, 11, 13 subfamilies, 4–11, 12 Genistein, 19, 20 Genomic structures, 4 Gentamicin, 166, 171 Geraniol, 182 Gingerol, 154 Glucose transporter 4, 17, 109, 322 Glutamate receptors, 18, 430 Glycine, 305 Glycoprotein, myelin-associated, 56 Glycosylation, 7, 107, 319 Golgi apparatus, 193, 319 G protein alpha subunits, 13 G-protein coupled receptor (GPCR) gene superfamily, 46 G-protein coupled receptors (GPCR), 19 blood vessels endothelial function, 379 vasoactive substances and, 363 Caenorhabditis elegans polycystin-1, 257 neuron pathﬁnding/migration, 59, 60 serine proteases, 422 TRPA1, 222 TRPC family activation mechanisms, 18, 21, 22, 36 protein–protein interactions, 335 TRPC signal transduction, 14 TRPM3, 147 TRPM5 function, 208 TRPP2 and PKD1, 196 TRPP2-related channels, 191 TRP-PLC signaling pathway, 15 TRPV2 and, 106–107, 112 vomeronasal receptors and, 46

Index G-protein coupled receptors, protease-activated receptors, 426 G-protein receptor kinases (GRKs), 426 Gqα, 13 Gq-coupled GPCR pathway, 14, 18, 19, 21, 22 Gq-PLC pathway, 2 Grasshopper chordotonal organ, 236–237 GRC, 276 Growth cone, neurons, 36, 57–58; see also Axon pathﬁnding, TRP channels and Growth factors, 34, 326 neurons, see Brain-derived neurotrophic factor (BDNF) TRPC5, 36 TRPV2, 18, 20, 105–106 Gt-PDE, 2 Guanine nucleotide exchange factor (GEF), 18 Guanosine triphosphate GTP-binding proteins TRPC protein–protein interactions, 339 TRPV4 regulation, 130 TRPC signal transduction, 14 Gustation/taste, 4, 234 TRPM4 function, 209 TRPM5, 4, 208, 278 Type I sensory neurons, 217 Gustatory nerve, 278 Gustducin, 13, 208

H Hair cells, 4, 222, 235 TRPA1, 164, 165, 166–167, 223 TRPV4, 129 Hair follicles, TRPV1 localization, 94 Hearing, see Auditory signals/hearing Heart TRPC2, 48 TRPV1, 94 TRPV2, 107, 389–394 TRPV4, 113, 126 Heart rate, anandamide triphasic response, 155 Heat activation, see Temperature Heat hyperalgesia, botulinum toxin and, 327 Heat receptors, thermoTRP channel gating biophysics, 288–289 Heat sensors, TRPV1 activation, 74 HEK293 cells, 19, 136, 142, 143, 264, 307, 308, 326 TRPC channel expression, 32 TRPM3 expression in, 145, 146–147 TRPV1 activation, 74 TRPV1 expression, 273 TRPV4 expression, 128, 129

449 HEK cells, 19, 20 protease-activated receptors, 431 TRPC family activation mechanisms, 22, 23, 24 Hemorrhagic cystitis, 155 Heteromeric receptors, 3, 356 protein–protein interactions, 332, 333; see also Protein–protein interactions, TRPC complexes TRPC family, 32, 35, 59, 60, 332, 333–334, 364 TRPV1/4, 309 vascular smooth muscle, 365 Hinging movements, channels, 120 Hippocampus, 326, 327 Histamine, 75, 363 Homer, 336, 341 Hormones, TRPV1 modulation, 95 Horseradish, 279 HPETE, 70, 75, 77, 88 Human autosomal polycystic kidney disease, see Polycystic kidney disease Human expressed sequence tags, see Expression sequence tags (ESTs) Humans, vomeronasal organs, 49–50 Hydrogen ions, see Protons/hydrogen ions/acids/pH Hydrophobic domains, 9, 12, 31–32 Hydrophobicity plots, 7 Hydroxycitronellal, 182 Hydroxyeicosatetraenoic acids, 73, 75, 77, 306 Hyperalgesia cold, 4 inﬂammatory, translocation of channels, 327 mechanical, 306, 430 protease-activated receptors, 432 thermal, see Thermal hyperalgesia TRPA1, 152–155 TRPV1 and, 76, 90, 96, 97 TRPV4 and, 118, 308–309 Hypernatremia, 405–406 Hypertension, 385, 386 Hypotension, cannabinoids and, 156 Hypothalamus main olfactory epithelium inputs, 50 protease signaling in nervous system, 428 TRPM3, 146 TRPV4, 132, 133 Hypotonic stress, see Osmoregulation/osmotransduction Hypoxia, 371

I IAV, see Inactive (iav) Icilin (AG-3-5), 170, 181, 182, 183, 184, 289

450

TRP Ion Channel Function in Sensory Transduction

binding sites, 352 TRPA1 sensitivity, 170, 171 Icrac, 3, 17, 35 IFT88, 232 Immunophilins, 336 Inactive (iav), 233, 304, 313 hearing and proprioception, 234–237 mechanotransduction, 222 INAD, 15, 335, 338 INDO, 381 Inﬂammation/inﬂammatory mediators protease-activated receptors, see Proteaseactivated receptors (PARs) temperature sensitivity, 288 translocation of channels, 327 TRPA1, 151, 158, 169 TRPM8, 279 TRPV1, 75, 76, 87–89, 90, 97, 273–274 TRPV4, 277, 308–309 Inner ear, 4, 163, 164–166, 171, 222, 223 Inositol triphosphate (IP3) endothelial function, 378 neuron pathﬁnding/migration, 59 signal transduction, 2 TRPC signal transduction, 14, 15–17, 21, 22, 33, 36 vasoactive substances and, 363 Inositol triphosphate (IP3) receptor genomic structures, 8, 12 neuron pathﬁnding/migration, 59, 60 TRPC2 binding, 48–49 TRPC protein–protein interactions, 339, 340–341 Insulin-like growth factor, 18, 276 channel translocation, 326 TRPV2 regulation, 105–112 calcium entry requirement for cell-cycle progression, 105–106 molecular identiﬁcation of channel, 107 property of channel, 106–107 regulatory process, 107–109 translocation of receptor, regulation of, 109–112 Insulin release, TRPV1 modulation, 95 Interleukins, TRPM4 function, 208 Intestinal CaT, 3 Intracellular compartments dissimilar structure-activity relationships, 92 translocation of receptors, see Trafﬁcking/translocation/sorting TRPM8, 185 TRPV1 modulation, 92, 95 TRPV2 localization, 107 Intraﬂagellar transport, 232, 260

Intrinsically photosensitive retinal ganglion cells (ipRGC), 4 Intrinsic modality elements, OCR-2, 249–251 Intron-exon boundaries, 9, 10, 11 Invertebrates, see also Caenorhabditis elegans; Drosophila osmo- and mechanotransduction, channel functions in, 303 thermonociception, 280–281 TRPA1, 171 TRPA1 in, 163, 164, 172–173 TRPP2 channels, 198 In vitro studies, see Cell culture In vivo studies, TRPV1 structure-activity relationships, 91–92 Inward calcium release-activated calcium current (Icrac), 3, 17, 35 Iodo-RTX, 93 Ion channels amino acid alignment, 7 permeability, see Permeability TRP channels in neuron pathﬁnding/migration, 56–57 Ionic conditions, see Osmoregulation/osmotransduction Ion selectivity, see Selectivity IP3, see Inositol triphosphate (IP3) Irritants, TRPA1 activation, 222 Isothiocyanates, 279, 280, 289

J Janus kinase 2, 191 Jasplakinolide, 17 Johnston’s organs, 229 Junctate, 336, 341

K KB-R7943, 24 KcsA potassium channel, 114, 115, 119, 120, 350, 352 Keratinocytes, 288 TRPV1, 94 TRPV4, 118, 132, 143, 277 Kidney renal ECaC discovery, 3 TRPM3, 145, 146 TRPP2 and polycystic kidney disease, 189–199; see also Polycystic kidney disease TRPV2, 107 TRPV4, 113, 117, 126, 127, 129, 143, 398–409 activation, 398–400 cellular osmoregulation, 400–401 systemic osmoregulation, 401–409

Index KIN-10, 258 Kinesin II mutants, 233, 260 Kinesin KLP-6 regulation, 259–260 Kinocilium, 229 KirBac1.1, 114 KLOV-1/PC-1, 259 KLP-6, 233, 258, 259–260 Knockdown models neuron pathﬁnding/migration, 58 TRPA1, 218, 223 Knockout models cannabinoid receptors, 155, 156 TRPC2, 49 TRPC4, 370, 371, 379 TRPC6, 385 TRPC family, 332 TRPM4/TRPM5, 208 TRPV1, 97, 306 TRPV4, 117, 143, 380

L Lamellopodia, 193 Lanthanum, 143, 190, 365 Latrunculin A, 110 Learning, olfactory, 245 Light perception, 234 Linalool, 182 Lingual nerve, 180 Lipid kinases, TRPM4/TRPM5 regulation, 207–208 Lipids, 234 thermoTRP channel gating biophysics, 294–295 TRPC signal complex organization, 337–338 trafﬁcking, 339 TRPV1, 71, 77 TRPV4 activation mechanisms, 398 gating mechanisms, 119, 120–121 Lipoxygenase anandamide metabolites in TRPV4 activation, 119 PLAT (polycystin/lipoxygenase/alpha toxin) domain, 191, 257–258, 260–261 TRPV1 pathways, 70, 71, 77, 88 Lipoxygenase homology 2 (LH2), 191 Lissencephaly 1 homology motif, 231 Liver TRPC2, 48 TRPV4, 126, 143 Localization

451 Caenorhabditis elegans TRPP, 258, 259–260 intracellular compartment translocation, see Trafﬁcking/translocation/sorting tissue expression, see Tissue expression/localization/distribution Locus coeruleus, TRPM3, 146 LOV-1, 196, 258, 259, 261, 262 Lung TRPV2, 107 TRPV4, 113, 126, 143 Lymphocytes NFAT (nuclear factor of activated T cells), 191, 192, 261 ROCE and SOCE, 23 Lyn, 130

M Macrophages, 107 Madin-Darby canine kidney cells, 196 MAG, 58 Magnesium, 147 Main olfactory bulb (MOB), 46, 50 Main olfactory epithelium (MOE), 46, 50 Male-speciﬁc sensory neurons, 196 MAP7, 115, 116, 129, 144 MAP kinases, 426 Mastalgia, 97 Mast cells, 91, 94, 106, 424, 429 MathI, 228 MCOLN3, 164 MDCK cells, 196 Mechanical hyperalgesia, 306, 430 Mechanical properties, TRPA1 molecular spring, 163 Mechanosensitivity/mechanotransduction, 280 Caenorhabditis elegans OCR-2 and OSM-9, see Caenorhabditis elegans cardiovascular system, 372, 389–394 endothelium, TRPV4 role in, 377–386 smooth muscle, 368 channel functions in, 303–314 C. elegans osm-9 gene, 310–313 future directions, 313 heterologous cellular expression system, 304–305 TRPV1, animal ﬁndings, 305–306 TRPV2, tissue culture data, 306–307 TRPV4, tissue culture and animal data, 307–310 TRPV subfamily, 304 Drosophila gene products in, 231–233 proprioception, 227, 228, 229–233

452

TRP Ion Channel Function in Sensory Transduction

Drosophila nociception innocuous versus noxious touch detection, distinct pathways, 216–217 mutants increasing thermal and mechanical nociception, 214–216 TRP channels, 222–223 endothelial function, 378, 379 neuron pathﬁnding/migration, possible roles, 61–63 osmotic changes and, 127 polycystins and, 259, 260 signal transduction, 312 TRPA, in Drosophila, 214–217, 222–223 TRPA1, 163, 164, 165, 171 discovery of, 3 hair cell transducer, 166–169 TRPP2, 196–199 TRPV (subfamily), 304 TRPV1, 305–306 TRPV2, 112, 306–307, 389–394 TRPV4, 113, 129–130, 143, 147, 377–386, 398–399 channel functions, 307–310 gating, molecular mechanisms, 116–117 interaction of different stimuli, 135 mechanism of, 130–131 Type I sensory neurons, 217 MEC proteins, 311 Melastatin subfamily (TRPM), see TRPM (subfamily) Membrane-spanning core, see Transmembrane domain Menthol, 7, 280, 288–289 Menthol receptor, 177–186, 217 METE ethanolamides, TRPV1 activation, 70 Methanandamide, 119 N-Methyl-D-glucamine (NMDG), 135 Methyl isothiocyanate, 170 Methyl salicylate, 154, 170 MHC class Ib, vomeronasal receptors and, 46 Microﬁlament-associated protein 7 (MAP7), 115, 116, 129, 144 Microtubules Drosophila mutants, 231–232 KLP-6 and, 259 neurons, cultured, 62 polycystins and, 259 Microvilli, sensory, 48 Mitochondria, 260 Mitogen-activated protein kinase, 192 Mitral cells, 46 Molecular spring, TRPA1, 163 Mouse embryonic ﬁbroblasts, 20, 22 Mucolipidin subfamily (TRPML), see TRPML Multidendritic sensory neurons, 215, 216, 218

Multiple irritant sensor, TRPA1 as, 159 Muscarinic receptor-stimulated calcium inﬂux, 18, 22 Mustard oils, 222, 279, 289 TRPA1, 152–155, 156, 159 activation by garlic derivatives and hazardous unsaturated aldehydes, 154–155 insensitivity to, 170 mediation of mustard oil effects in sensory neurons, 153–154 Myelin-associated glycoprotein (MAG), 56 Myocytes, TRPV2, 112, 390–394 MyoVIIA, 233 Myristoylation, TRPV4, 114, 115

N NADA (N-arachidonoyldopamine), 76, 88 Nanchung, 228, 233, 304, 313 Drosophila, hearing and proprioception, 234–237 mechanotransduction, 222 NAPE (N-arachidonoylphosphatidylethanolamine), 70, 73 N-arachidonoyldopamine (NADA), 76, 88 N-arachidonoyl-phosphatidylethanolamine (NAPE), 70, 73 N-arylcinnamides, 93 NCX (sodium-calcium exchanger), 23, 24, 342 Nematodes, TRPA1, 171 Nerve growth factor (NGF), 90, 97, 158, 183, 326, 430, 431 Nervous system, see also Brain; Central nervous system; Neurons axon pathﬁnding, see Axon pathﬁnding, TRP channels and protease-activated receptors (PARs), 427–433 enzymes, 429 expression in, 427–429 regulation of neuronal excitability, 429–430 sensitization of TRPV channels, 430–433 TRPA1 in nerve injury, 158 Netrin, 4, 56, 57, 58, 61 Neurite outgrowth, 18 Neuroendocrine cells, TRPV2, 107, 112 Neuroendocrine factors, TRPV1 regulation, 95 Neuroﬁlaments, cultured neurons, 62 Neurogenic inﬂammation ethanol and, 88–89 protease-activated receptors, see Proteaseactivated receptors (PARs) translocation of channels, 327 TRPA1 and, 151 TRPV1 regulation, 274

Index Neurokinin (NK1) receptors, mast cells, 91 Neurons axon pathﬁnding, TRP channels and, 55–63 mechanosensation in migrating neurons, 61–63 neuronal migration guidance factors, 61 thermosensation for migration, 63 growth cones, 36, 57–58 LOV-1 and PKD-2, 259 neurite extension, 18 pain pathway probes, 152–155 PAR regulation of excitability, 429–430 temperature sensitivity, 288 thermosensory, functions of high-threshold and low-threshold types, 221 TRPA1, 152–155, 163 TRPC2 and pheromone detection in mammals, 46 TRPC5 protein–protein interactions, 339 TRPM8, 177–186 TRPP2 and PKD1 coexpression, 195 TRPV1 activation pathway, 70–72 coexpression and overlapping activity, 96 localization, 95 TRPV2, 107, 112 TRPV4, 126 Neuropathic pain, TRPA1, 169 Neurotransmitters and neuron growth cone calcium, 56 protease signaling in nervous system, 428 TRPV1 modulation, 90 Nexin 1, 426, 427 NFAT (nuclear factor of activated T cells), 191, 192, 261 NHERF, 15, 335, 336, 337, 339 Nickel, 106 Nitric oxide/nitric oxide synthase, 14, 370, 371, 378, 381, 383, 384 NMDA receptors, 61, 305, 430 NMDG (N-methyl-D-glucamine), 135 Nociception/pain pathways, 234 Caenorhabditis elegans, 249 Drosophila, 213–223; see also Drosophila, nociception pain perception/signal transmission, 69–70 protease-activated receptors, see Proteaseactivated receptors (PARs) thermosensation, see Thermosensation TRPA1, 151, 159, 170–171 activation by garlic derivatives and hazardous unsaturated aldehydes, 154–155 channel properties, 172 function in, 170–171 mediation of mustard oil effects in sensory neurons, 153–154

453 mustard oil and capsaicin as chemical probes of, 152–155 mustard oil and capsaicin as chemical probes of pain pathway, 156 sensory ganglia, 169 sensory ganglia, distribution in, 169 TRPM8, cold sensing and menthol receptor, 180, 181 TRPV1, 87–89 TRPV4, 4, 147 mechanisms of, 144 molecular mechanisms of gating, 117 Nodose ganglion, TRPA1, 163, 169 NompA, 216, 232, 235; see also TRPN (subfamily) NompB, 232 NompC, 164, 198, 216, 222 Norepinephrine, 363 NorpA Drosophila mutant, 15 N-type calcium channel, cultured neurons, 62 Nuclear factor of activated T cells (NFAT), 191, 192, 261 Nucleotides, 363 NUDIX domain, 4, 7, 10, 12 NUDT9, 351, 355

O OAG (oleyl-acetyl-glyceride), 20–21, 34 OCR-2, 260, 304, 310–311; see also Caenorhabditis elegans OKD-2/PC2, KLP-6 colocalization, 259 Oleoylethanolamide (OEA), 76 Oleyl-acetyl-glyceride (OAG), 20–21, 34 Olfaction/olfactory signals, 45–51; see also TRPC2, and pheromone detection in mammals Caenorhabditis elegans, learning in, 245 signal transduction, 312 TRPV4, 380 Type I sensory neurons, 217 Olfactory bulb, 429 Oligomerization of cytosolic domains, 356 Olvanil, 72 Open reading frames (ORFs), 7–8, 9, 12, 13 Organ of Corti, 164, 165 Organosulfur compounds, 153–155, 156; see also Mustard oils OSEG mutants, 232 OSM-5, 232 OSM-9, see Caenorhabditis elegans, OSM-9 Osmoregulation/osmotransduction, 7, 280, 306 Caenorhabditis elegans, 249 channel functions in, 303–314 Drosophila nan and iav, 234

454

TRP Ion Channel Function in Sensory Transduction

protease-activated receptors, 432–433 signal transduction, 312 temperature sensitivity, 288 TRPM3, 4, 145–147 TRPM4, 4 TRPV2, cardiovascular system, 391–392 TRPV4, 113, 114, 117, 126, 127–129, 380, 397–409 calcium and, 134 interaction of different stimuli, 135 mechanism of regulation, 130–131 TRPV4 activation by hypotonicity, 399–400 TRPV4 in cellular osmoregulation, 400–401 TRPV4 in systemic osmoregulation, 401–409 anatomy of kidney, 403–404 model of physiological function, 404–405 water balance in vivo, 405–409 Osmoregulation/osmotransduction, volume regulation, see Volume regulation OTRPC4, see TRPV4 Outer segment genes, 232 Overlapping activity, TRPV1, 96–97 Oxidative stress, endothelial cell response, 372

P PACS-1 and PACS-2, see Phosphofurin acidic cluster proteins (PACS-1 and PACS-2) painless gene, 171, 280–281; see also Drosophila, nociception Pain pathways, see Nociception/pain pathways Pancreas TRPM4, 203, 204 TRPV1 modulation in islet cells, 95 TRPV2, 107 Parasites, TRPA1, 171 PARs, see Protease-activated receptors (PARs) Pathological conditions, changes of TRPV1 expression, 97 PC-1, 261 PDD-4α, see Phorbol didecanoate-4α (PDD-4α) PDZ domains, 14–15, 321, 335, 337 Pericentrin-like protein, 231 Pericentriolar material, 231 Permeability TRPM4/TRPM5, 204–205 TRPV4, 119–120 vascular capillaries, 371 endothelium, 362 Pertussis toxin, 106–107 pH, see Protons/hydrogen ions/acids/pH Pharmacokinetics, TRPV1 structure-activity relationships, 91–92 Pharmacological properties

overlap between TRPV1 and its relatives, 96–97 TRPM3, 146–147 TRPV4, 143–144 Phenylephrine, 364, 365 Phenymethylsulfonyl ﬂuoride, 119 Pheromones, 45–51; see also TRPC2, and pheromone detection in mammals Phorbol didecanoate-4α (PDD-4α), 119, 131, 132, 134, 280, 308, 398 dissimilar structure-activity relationships, 119 endothelial function, 378, 380, 382–383, 386 interaction of different stimuli, 135 protease-activated receptors, 433 Phorbol esters, 7 capsaicin analogues, 73 endothelial function, 378, 380 translocation of channels, 327, 328 TRPV1, 76 TRPV4, 113–114, 128, 144, 398 activators, 131–132 calcium and, 134 molecular mechanisms of gating, 118 Phorbol myristate acetate (PMA), 76, 128, 131, 398 Phosphatases, see also Calcineurin TRP-PLC signaling pathway, 15 TRPV1 modulation, 89, 91 Phosphatidylinositol 3-kinase (PI3K), 192 channel translocation, 326 protease-activated receptors, 426 TRPV1 modulation, 90–91 TRPV2, 106, 110 Phosphatidylinositol 4,5-bisphosphate (PIP2), 2, 7 pharmacological overlap, 96 pore domain structure, 352 protease-activated receptors, 426, 431 receptor structure, 321 TRPA1, 158 TRPC family, 18 activation mechanisms, 31, 33, 36 signal transduction, 14 trafﬁcking, 339 TRPM4/TRPM5, 207–208, 210 TRPM8, 183–184, 279 TRPV1, 77, 89–90, 96 vasoactive substances and, 363 Phosphatidylinositol 4-kinase, 34 Phosphatidylinositol-4 phosphate 5-kinase (PIP5K), 18, 326 Phosphodiesterases, TRPV1 activation pathway, 70, 71 Phosphofurin acidic cluster proteins (PACS-1 and PACS-2), 142, 144, 190, 193 Phosphoinositide 3-kinases, 34

Index Phospholipase A2 ciliated epithelia, 417 endothelial function, 378 TRPV1, 77 TRPV4, 117, 400, 417 and vasodilatation, 384 Phospholipase C (PLC), 191, 280 blood vessels smooth muscle cells, 364 vasoactive substances and, 363 neuron pathﬁnding/migration, 59 translocation as activation and regulatory mechanism, 324 TRPA1, 158, 170 TRPC family activation mechanisms, 14–15, 19, 31, 33, 35, 36 TRPM4/TRPM5, 208, 209 TRPM8, 183–184, 279 TRPV1, 90 TRPV4, 116–117, 118 Phospholipase Cβ (PLCβ), 14, 21, 426, 430, 432 Phospholipase Cβ2 (PLCβ2), 208 Phospholipase Cγ (PLCγ), 14, 20–21, 60, 337–338, 431 Phospholipase Cγ1 (PLCγ1), 337–338 Phospholipids, TRPV1 activation pathway, 70 Phosphorylation/dephosphorylation, see also speciﬁc kinases and phosphorylases insulin-like growth factor binding and, 106 TRPC signal transduction, 14 TRPP2 channel modulation, 199 TRPV2, 107 TRPV4 structure, 114 Phosphotyrosine, cultured neurons, 62 Phospho-Y226, 19 Photoreceptor cells, 33, 364, 379 dynamic translocation of, 325 protein–protein interactions, 335 Phototransduction, 2, 4, 46 Phylogenetic comparison, 4, 7 temperature sensitive TRP channels, 272 TRPA1, 164 PI3K, see Phosphatidylinositol 3-kinase (PI3K) PIGEA-14, 193 PIP2, see Phosphatidylinositol 4,5-bisphosphate (PIP2) Piperine, TRPV1 activation, 73 PKD1, 199, 257 PKD1-like proteins, 190–191 PKD1L13, 190 PKD1-TRPP2 complex activation of, 195–196 cellular effects, multiple functions, 194–195 PKD2, 144, 190, 257, 258, 260, 262; see also TRPP2 (polycystin-2)

455 PKD2-like proteins, 190 PKD-REJ (receptor for egg jelly, TRPP4), 6, 11, 190, 191, 195, 196 Placenta TRPM4, 204 TRPV4, 126 Plasma membrane, channel trafﬁcking, see Trafﬁcking/translocation/sorting Plasminogen N-terminal (PAN) modules, NompA, 232 PLAT (polycystin/lipoxygenase/alpha toxin) domain, 257–258, 260–261 PLAT/LH2, 191 PLC, see Phospholipase C (PLC) PLIK kinase domain, 12, 321 PMD38, 182 Polaris IFT complex B polypeptide, 260 Polaris/Tg737, 232 Polarization, neurons in growth cones, 56 Polycystic kidney disease Caenorhabditis elegans, polycystin ciliary localization and ADPKD, 257–259 PKD1-TRPP2 complex activation of, 195–196 cellular effects, multiple functions, 194–195 TRPP2, activation mechanisms and functional roles, 189–199 mutations causing, 191–193 Polycystin-1, 257–259 Polycystin-2, see TRPP2 (polycystin-2) Polycystin channel protein PKD2, 144 Polycystin subfamily (TRPP), see TRPP (subfamily) Poly-L-lysine, 207 Polymerase chain reaction (PCR), TRPV2, 109 Polyneuropathies, TRPV1 and, 96 Pore loop/pore region, 8, 9 gating mechanisms, 120 protein–protein interactions, see Protein–protein interactions, TRPC complexes sequence similarity among TRP family, 142 structure, 350–352 TRPV1, 70 TRPV4, 114, 115–116, 119–120, 142 Postranslational modiﬁcation, 319 phosphorylation/dephosphorylation, see speciﬁc kinases and phosphorylases protease-activated receptors, 425 TRPV1, 87; see also Splice variants TRPV2, 107 TRPV4, 114, 115 Potassium channels endothelial calcium-activated, 378, 383 KcsA, 114, 350

456

TRP Ion Channel Function in Sensory Transduction

MlotiK1, 356 sensor domain, 352 Shaker, 250–251, 294 thermosensory, 219 TRPC2 and, 48 TRPM4 function, 208–209 TRPP2-related channels, 190 TRPV4, 308 TRPV4 analogy, 115 PP2, 20 Pressure-induced changes blood vessels, 363 vascular function, smooth muscle cells, 362 vascular smooth muscle, 365 vascular smooth muscle cells, 366–368 vasoconstriction agonist-induced channel activation, 363–366 vasodilator response to EETs mediated by channel activation, 366 Primates pheromone sensing, molecular biology of, 50 vomeronasal system, 47 Progression factors, cell cycle, 105–106; see also Insulin-like growth factor, TRPV2 regulation Proline rich domain, 336, 340 2-Propenal, 155 Proprioception Drosophila, see Drosophila, hearing and proprioception mechanosensitivity and, 303 Prostacyclin, 378 Prostaglandins, 382 endothelial function, 378 osmo- and mechanotransduction, channel functions in, 309 protease-activated receptors, 426 TRPV1 regulation, 75, 90, 274 vascular endothelium, 370 Prostate cancer cells, TRPV1 and TRPM colocalization, 96–97 Prostate gland, 189, 185, 204 Protease-activated receptors (PARs), 91, 274, 370, 371, 422–434 cleavage and activation, 422–426 enzymes, 423–426 molecular mechanisms of activation, 422–423 nervous system, 427–433 enzymes, 429 expression in, 427–429 regulation of neuronal excitability, 429–430 sensitization of TRPV channels, 430–433 signal transduction mechanisms, 426–427

Proteinase C, 426 Protein denaturation, thermoTRP channel gating biophysics, 294–295 Protein kinase A (PKA), 191, 192 protease-activated receptors, 426, 432 TRPV1, 75, 76, 89, 90–91 activators, 70 heat-induced, 74 intracellular compartment translocation, 91 TRPV2, 107 Protein kinase alpha domain, 4 Protein kinase C (PKC), 191, 192 neuron pathﬁnding/migration, 59 protease-activated receptors, 432 signal transduction, 2 translocation of channels, 327, 328 TRPC family activation mechanisms, 17, 22, 36 TRPM4, endothelial, 380 TRPM8 regulation, 184 TRPV1, 75, 76, 77, 89, 90–91, 275 activators, 70 dissimilar structure-activity relationships, 92 heat-induced, 74 intracellular compartment translocation, 91 mast cells, 91 TRPV2, 107 TRPV4, 113–114, 118 phorbol ester effects, 132 structure, 115 Protein kinase Cε (PKCε), 76, 432 Protein kinase CK2 (PKCK2), 260, 261 Protein kinase D1 (PKD1), 432 Protein kinases protease-activated receptors, 426 TRPC signal transduction, 14 TRPV1 modulation, 90–91 TRPV4, 144–145 Protein–protein interactions ankyrin repeats and, 163, 353 channel trafﬁcking, 320, 321 oligomerization of cytosolic domains, 356 TRPC family activation mechanisms, 15–17 Protein–protein interactions, TRPC complexes, 332–343 abbreviations, 343 anchoring of channels in signalplexes, 335–338 cellular trafﬁcking of channel complexes, 338–339 gating, 339–341 activation/deactivation, potential interactions involved in, 340–341 inactivation, potential interactions involved in, 341–342 oligomerization and assembly of pore complexes, 333–335

Index signaling partnerships, 342 subunit assembly and anchoring of channels, 333–338 versatility of receptor components, 332–333 Protein sorting, see Trafﬁcking/translocation/sorting Protein trafﬁcking, see Trafﬁcking/translocation/sorting Protonation, TRPV1 receptor structure, 70 Proton hopping, 74, 88 Protons/hydrogen ions/acids/pH, 7, 280, 288, 324 proteases and, 430 sodium-proton exchanger, 15, 335, 336, 337, 339 TRPM4/TRPM5, 205 TRPM5, 204 TRPM8, 184 TRPP2, 199 TRPV1, 73–74, 76, 88, 273, 275 TRPV4, 113 Pseudogenes, 32, 50 Purine nucleotides, 363 Purkinje cells, TRPV2, 107 pyrexia mutants, 218, 219, 220 Pyrimidine nucleotides, 363

R Rac1, 18, 34, 326 raf-1, 426 Randall-Sellito test, 308 Rapamycin, 15 Rapid vesicular insertion of TRP (RiVIT), 326 Rapid vesicular trafﬁcking, TRPC, 339 Ras, 106 Receptor for egg jelly, see PKD-REJ (receptor for egg jelly, TRPP4) Receptor-operated calcium entry (ROCE), 2–3, 19 ROCE SOCE conundrum, 2–3, 19 TRPC family activation mechanisms, 21, 22, 23–24 Receptor operated channel operating with TRPV1, TRPA1, 158–159 Rectum, TRPV1, 305–306 Reducing agents, TRPV1, 77–78 Regulatory volume decrease (RVD), 307, 415 Renal ECaC, 3 Reproduction, mating, fertilization Caenorhabditis elegans, 257–264, 325 TRPC2 and, 45–51; see also TRPC2, and pheromone detection in mammals Resiniferatoxin (RTX), TRPV1 activation, 73, 86, 87 pathological conditions, changes of expression under, 97 structure-activity relationships, 91–92, 93

457 Respiratory tract ciliated epithelium, see Cilia/ciliated epithelium protease-activated receptors, 433 TRPV1, 88 TRPV4, 126, 143, 308, 414 Retinal ganglion cells, 4 RhoA, 336, 339 Rho GTPase, 326 Rho kinase, 426 Rhyanodine channels, 61 Rimonabant, 155 ROCE, see Receptor-operated calcium entry (ROCE) Rsp defects, 259 RTK, 22 RTK-PLCg pathway, 21 Ruthenium red, 166, 367, 378, 382 TRPA1, 153, 155, 156, 171 TRPV4, 119, 130, 136 Ryanodrine receptors, 366

S Salivary glands, TRPV4, 126, 143 Sarcoplasmic reticulum, 363–364 SERCA (sarcoplasmic-endoplasmic reticulum calcium pumps), 2, 15–16 SB-452533, 96 Scaffolding protein, 34 protein–protein interactions, 337 TRPM3, 147 Scolopidium structure, Drosophila, 230–231 SDF, 56, 61 Sea urchin, 196 Sebocytes, TRPV1 localization, 94 Secretory epithelia, TRPV4, 308 Selectivity pore domain structure and, 350, 352 TRPV4, 119–120 Semaphorin, 58 Sensillium, 216, 217 Sensor domain structure, 352–353 Sensory ganglia temperature sensors in mammals, 217 TRPA1, 163 auditory organs, 169 nociception, 169 TRPV4, 113 SERCA (sarcoplasmic-endoplasmic reticulum calcium pumps), 2, 15–16 Serine proteases, 430; see also Protease-activated receptors (PARs) Serotonergic neurons, 243–252 Serotonin, 75, 313, 363

458

TRP Ion Channel Function in Sensory Transduction

Sex differences/dimorphism pheromone sensing, molecular biology of, 49, 50 pkd-2, 196 TRPV1 activation, 95 vomeronasal organ discrimination, 50 SFK (src family kinase), TRPV4, 130 Shaker potassium channels, 250–251, 294, 351, 352, 353, 356 Shear stress, see also Mechanosensitivity/mechanotransduction TRPP2, 196–197 TRPV2, 393 TRPV4, 113, 116, 143, 377–386 vascular endothelium, 377–386, 393 Shuttling, intracellular, see Trafﬁcking/translocation/sorting Signal ampliﬁcation, neuron guidance factors, 61 Signal transduction, see also speciﬁc channels and channel families Caenorhabditis elegans OCR-2 and OSM-9, 249 odorant, osmotic, and mechanical stimuli, 312 pheromone signaling, 46, 47, 48 Silkworm cell expression system, 2 Skeletal muscle, 15–16, 107 SKF-96365, 58, 365 Skin TRPV1, 94, 97 TRPV4, 118, 143, 277 Skin cancer, TRPM8, 185 Skin disease, TRPV1 in, 97 Slit, 56, 61 Small GTP binding proteins, TRPV4 regulation, 130 Small interference RNA (siRNA), 18 neuron pathﬁnding/migration, 58 TRPC family, 332 TRPC family activation mechanisms, 23 Smooth muscle, 17, 18 ROCE and SOCE, 23 TRPM8, 185 TRPP2, 194 TRPV1, 94 TRPV2, 112 TRPV4, 143, 380–381 vascular function, 363–369 pressure-induced channel activation, 366–368 store-operated channel activation, 368–369 TRP channels, 362 vasoconstriction agonist-induced channel activation, 363–366 vasodilator response to EETs mediated by channel activation, 366

Snail neuron growth cones, 62 SNAP, 339 Snapin, 327 SNARES (soluble N-ethylmaleimide-sensitive factor attachment protein receptors), 320–321, 325, 326, 327, 328, 339, 354 SOCE, see Store-operated calcium entry (SOCE) SOCs, see Store-operated calcium channels (SOCs) Sodium, TRPV4 in systemic osmoregulation, 405–406 Sodium-calcium exchanger, 23, 24, 342 Sodium channels, channel-activating proteases, 430 Sodium-proton exchanger, 15, 335, 336, 337, 339 Sodium pump, 23 Soluble N-ethylmaleimide-sensitive factor attachment protein receptors (SNAREs), 320–321, 325, 326, 327, 328, 339, 354 Somatosensory ganglia, TRPA1, 164 S100, 327 Sorting, protein, see Trafﬁcking/translocation/sorting spe-9, spe-38, spe-42, 264 Sperm, 196 TRP-3 in Caenorhabditis elegans mating and fertilization, 32, 262–264 TRPC2, 48, 49 Sphingosine, 146 Spinal neurons, neuron pathﬁnding/migration, 58 Spleen, 48, 107 Splice variants, 4 TRPC2, 48, 49 TRPC2β, 48 TRPM3, 146 TRPM4, 203 TRPV1, 75 possible regulation by, 95 tissue and cellular expression, 94 TRPV4, 142, 143 Spodoptera frugiperda, 2 Spontaneous transient outward currents (STOCs), 366 Spring, molecular, 163 SR141716A, 155 src kinases, 19, 21 TRPV1, 91 TRPV4, 114, 130, 145 Stathmin domain, 338–339 Stereocilia, 165, 228, 229 Steroids, TRPV1 regulation, 95 STIM1, 2, 17, 33 TRPC family activation mechanisms, 21, 36, 37 TRPC protein–protein interactions, 341 TRPC signal transduction, 14

Index STOCs (spontaneous transient outward currents), 366 Stomach TRPM5, 204 TRPM8, 185 TRPV1, 94 Store depletion, 35 Store-operated calcium channels (SOCs), 33, 34, 323, 340 trafﬁcking of channel complexes, 323, 324–325 vascular function endothelium, 369 smooth muscle, 362, 368–369 Store-operated calcium entry (SOCE), 2–3, 19, 32 candidates for channel, 33 and TRPC family, 23–24 TRPC family, 36 activation mechanisms, 21, 22 protein–protein interactions, 341–342 vascular endothelium, 370–371 Stretch activation endothelial function, 379 and neuron migration, 62 TRPV2 and, 112 Type I sensory neurons, 217 vascular smooth muscle, 362 Strontium, 133 Structure, see also speciﬁc features (ankyrin repeats, transmembrane region, etc.), 349–356 amino acid sequence alignment, 7, 8 cytosolic domains, 353–356 ankyrin repeats, 353–355 enzymatic domains, 355 MHR in TRPM channels, 355 oligomerization of, 356 TRPML and TRPP subfamilies, 355–356 OCR-2, 250–251 polycystin-1, 257–258 polycystin-2, 258–259 protein–protein interactions, see Protein–protein interactions, TRPC complexes scaffold, 349–350 transmembrane region, 350–353 pore domain, 350–352 sensor domain, 352–353 TRPM4/TRPM5, 204 TRP receptor subunit, 321 TRPV1 structure-activity relationships capsaicin sensitivity, 86–87 dissimilarity of, 91–92, 93 Subcellular localization, see Trafﬁcking/translocation/sorting Substance P, 279, 428, 429

459 Substantia nigra, 126, 146 Sulfhydryl groups, TRPV1 modulation, 77–78 Sulfur compounds, TRPA1 activation, 152–155, 156 Suprachiasmatic nucleus, 4 Sweat glands, TRPV1 localization, 94 SYF cells, 19 Synaptic vesicle proteins, 327 Synaptic vesicle recycling, 320 Synaptobrevin 2, 321 Synaptogamin IX, 327 Syntaxin 1, 321 Syntaxin 3, 339

T Tactile sensation, see Touch detection Taste, see Gustation/taste TAX-6, 258, 261 Taxol, 309 T cells, see T lymphocytes Tear glands TRPA1, 159 TRPV4, 308 Tectorin, 232 Temperature, 7, 234, 362; see also Thermosensation Drosophila, TRPA channels in nociception combinatorial encoding of temperature response, 218–220 multiple heat activated channels in ﬂies, 218 thermosensory neurons, functions of highthreshold and low-threshold types, 221 and osmotic sensitivity, 127 TRPA1 heat activation, 158–159 TRP channels in neuronal migration, possible roles, 63 TRPV1, 4 TRPV1 heat activation, 74–75, 87–88 ethanol and, 89 pathological conditions, changes of expression under, 97 PKC phosphorylation and, 76 sulfhydryl group oxidation state and, 78 TRPV2, 4, 393–394 TRPV3, 4 TRPV4, 113, 132–133, 144 activation by thermal stress, 398 interaction of different stimuli, 135 mechanosensitivity, 143 molecular mechanisms, 117–118 thermoregulation, 132, 133 vascular function, endothelial cell response, 371, 393–394

460

TRP Ion Channel Function in Sensory Transduction

Tetanus neurotoxin, 325, 326 Tetradecanoylphorbol-13-acetate (TPA), 327, 328 Tetrahydrocannabinol (THC), 155, 156, 280 Thapsigargin, 2, 16, 20, 325–326, 369 TRPC2 in sperm cells, 49 TRPC family activation mechanisms, 22, 23 Thermal hyperalgesia, 153; see also Thermosensation protease-activated receptor sensitization of TRPV channels, 430 TRPA1, 158, 218 TRPV4, 132–133 Thermosensation, 234, 271–281; see also Temperature channel gating biophysics, 287–299 allosteric model, 297–299 cold receptors, 289 heat receptors, 288–289 minimum model to explain temperatureinduced changes, 296–297 multistate nature of channel opening process, 295–296 protein denaturation and lipids and, 294–295 temperature dependence, 289–291 voltage dependence, 291–294 cold receptors, 278–280 channel gating biophysics, 289 TRPA1, 158, 222 TRPA1 (ANKTM1), 279–280 TRPM8 (CMR1), 177–186, 278–279 heat receptors, 273–276 channel gating biophysics, 288–289 TRPV1 (VR1), 273–275 TRPV2 (VRL-1), 275–276 invertebrate thermonociception thru TRP channels, 280–281 temperature sensors in mammals, 217–218 warm receptors, 276–278 TRPM4 and TRPM5, 277–278 TRPV3 and TRPV4, 132, 276–277 Thermosensory neurons combinatorial encoding of temperature response, 218–219 high-threshold and low-threshold types, 221 Thio-BCTC, TRPM8 antagonists, 183 Thiosulﬁnates, TRPA1 activation, 154 Thrombin, 371, 429, 430, 432 Tissue expression/localization/distribution TRPC2, 48 TRPM3, 145–146 TRPM4/TRPM5, 204 TRPV1, 73, 94 TRPV2, 107 TRPV4, 113, 126–127, 142–143

Tissue plasminogen activator (TPA), 430 T lymphocytes NFAT (nuclear factor of activated T cells), 191, 192, 261 ROCE and SOCE, 23 TRPM4 and, 208, 209 Tongue, TRPM5, 204 Touch detection, see also Mechanosensitivity/mechanotransduction Caenorhabditis elegans, 249 Type II sensory neurons in, 216 Type I sensory neurons, 217 Touch-insensitive larval B (TilB), 233 Trachea, TRPV4, 126, 143 Trafﬁcking/translocation/sorting, 319–328 as activation and regulatory mechanism, 324–327 agonist-induced translocation of TRPC channels, 325–326 functional modulation of TRPL and TRP3 channels, 325 modulation of TRPV channel activity, 326–327 translocation to plasma membrane, activation of SOC channels, 324–325 polycystin-2, 259 protease-activated receptors and, 427–428 protein–protein interactions, signal complex organization, 337 TRP-3 in Caenorhabditis elegans sperm activation, 263–264, 325 TRPC family, 338–339 activation mechanisms, 17–18, 35 binding partner effects on channel localization and function, 336 signal transduction, 14 TRPC5 channels, 36 TRPP2, 193 TRPV1, 91, 95 TRPV2, 108, 276 Tranilast, 106 Transducins, 13 Transferrin, 109 Transition state theory, 296–297 Translocation of receptors, see Trafﬁcking/translocation/sorting Transmembrane domain, 355 structure, 350–353 pore domain, 350–352 sensor domain, 352–353 TRPC family, 334–335 TRPM3, 147 TRPM8, 185 TRPP subfamily, 190 TRPV1, 70, 72

Index TRPV2, 107–108 TRPV4, 114, 115–116, 130, 142 Trapdoor model, 235, 236 Trichromatic vision, 46, 47, 50 Trigeminal ganglia temperature sensors in mammals, 217 TRPA1, 170 TRPM8, 178, 180 TRPV4, 113, 126 TrkA receptor, 158, 169, 180, 183–184 TrkB, 59 TRP (superfamily), 234 diversity of form and function, 2–4 genes and gene products, 4–13 neuron pathﬁnding/migration, 55–63 TRP-1, 262 TRP-2, 262 TRP-3, 262–264 TRP-4, 251 TRPA (subfamily), 323, 363 ankyrin repeats, 353, 354 Drosophila, nociception combinatorial encoding of temperature response, 218–220 innocuous versus noxious touch detection, distinct pathways, 216–217 multiple heat activated channels in ﬂies, 218 mutants increasing thermal and mechanical nociception, 214–216 temperature sensors, 272 temperature sensors in mammals, 217–218 thermosensory neurons, functions of high-threshold and low-threshold types, 221 genomic structures, 9 TRP subfamilies, 126, 190 TRPA1 (ANKTM1), 3, 4, 151–159, 198, 304 amino acid sequence alignment, 8 auditory organs, 164–169 channel properties of TRPA1 versus hair cell transducer, 166–167 inner ear, distribution in, 164–165 inner ear, functional tests in, 165–166 sensory ganglia, 169 cannabinoid activation, 155–157 cold sensation, 157–158 GenBank data, 6 genomic structures, 10 in invertebrates, 172–173 as multiple irritant sensor, 159 mustard oil and capsaicin as chemical probes of pain pathway, 152–155 activation by garlic derivatives and hazardous unsaturated aldehydes, 154–155, 156 mediation of mustard oil effects in sensory neurons, 153–154

461 nociception, 170–171 channel properties, 172 function in, 170–171 multiple elicitors of burning pain, 222 sensory ganglia, 169 receptor operated channel operating with TRPV1, 158–159 temperature sensitivity, 288, 289 thermosensation, 272 chemical agonists, 280, 281 painless gene, 280–281 role of TRP channels, 279–280 touch transduction, 235 TRPV1 coexpression, 96 TRP box, 323 TRPC (subfamily), 2–24, 234, 323, 363 activation mechanisms, 14–24, 31–37 conformational coupling, IP3 receptor role, 15–17 diacylglycerol and protein kinase C, 17 GPCR-Gq activation to voltage-gated calcium channel activation, 18 phospholipase C, 14–15 protein–protein interactions, channel gating, 339–341 in situ, 35–36 in transfecto, 32–35 translocation of channels to plasma membrane and calcium entry, 17–18 tyrosine kinases in voltage-gated calcium channel-independent calcium inﬂux, 19 blood vessels/vascular structures, 362, 364 Caenorhabditis elegans mating and fertilization, 262–264 questions and future directions, 264 translocation during sperm activation, 325 TRP-3 and sperm-egg interactions, 262–263 TRP-3 translocation during sperm activation, 263–264, 325 discovery of, 2–3 endothelial function, 378, 379 neuron pathﬁnding/migration, 58–59 nomenclature, 4 pore forming sequence similarity, 142 protein–protein interactions, see Protein–protein interactions, TRPC complexes SOCE and, 23–24 structure ankyrin repeats, 353, 354, 355 pore domain, 352 terminology, 3 TRP subfamilies, 125, 190

462

TRP Ion Channel Function in Sensory Transduction

TRP superfamily diversity, 2–4 TRP superfamily genes and gene products, 4–13 direct chromosomal repeats, 12–13 genomic structures, 11, 13 subfamilies, 4–11, 12 tyrosine phosphorylation, role in function of, 19–24 TRPC1, 192 blood vessels/vascular structures, 362, 364 endothelium, 369–371 smooth muscle, 365, 366, 368 store-operated channel activation, 368–369 endothelial function, 379 GenBank data, 5 heteromeric channels, 32, 333 mechanoregulation, 199 protein–protein interactions, 339 activation/deactivation, potential interactions involved in, 340–341 binding partner effects on channel localization and function, 336 IP3 receptor, 340 signal complex organization, 337 SOCE, 23 Xenopus spinal neuron pathﬁnding, 58 TRPC1/6 heteromeric channel, vascular smooth muscle, 365 TRPC2 blood vessels/vascular structures, 362, 364 GenBank data, 5 genomic structures, 11 protein–protein interactions activation/deactivation, potential interactions involved in, 341 binding partner effects on channel localization and function, 336 TRPC2, and pheromone detection in mammals, 4, 45–51, 263 molecular biology of pheromone sensing, 46–48 protein structure and binding partners, 48–51 evolution of pheromone sensing, 49–50 mechanism of activation, 49 from vomeronasal activation to behavioral changes, 50–51 TRPC3, 7, 24 amino acid sequence alignment, 8 blood vessels/vascular structures, 362, 364 endothelium, 370, 372 smooth muscle, 365, 366 endothelial function, 379 GenBank data, 5 heteromeric channels, 32, 59, 60, 333

neuron pathﬁnding/migration, 4, 59 protein–protein interactions activation/deactivation, potential interactions involved in, 340, 341 binding partner effects on channel localization and function, 336 signal complex organization, 337 signaling partnerships, 342 SNARE complexes, 339 signal transduction, 17 SOCE, 23 translocation of, agonist-induced, 325–326 tyrosine kinases in voltage-gated calcium channel-independent calcium inﬂux, 19 tyrosine phosphorylation, role in function of, 19 TRPC3/6 heteromeric channels, 60 TRPC4, 5, 15, 263 blood vessels/vascular structures, 362, 364 endothelial function, 370–371, 379 heteromeric channels, 32, 333 protein–protein interactions binding partner effects on channel localization and function, 336 channel anchoring, 335 signal complex organization, 337 signal transduction, 17 SOCE, 23 TRPC5, 24 activation mechanisms, 15, 17 blood vessels/vascular structures, 362, 364 GenBank data, 5 genomic structures, 11 heteromeric channels, 32, 333 neurons in growth cones, 57–58 protein–protein interactions activation/deactivation, potential interactions involved in, 341 binding partner effects on channel localization and function, 336 channel anchoring, 335 trafﬁcking of channel complexes, 338–339 translocation of, agonist-induced, 325–326, 327 TRPC6, 24, 263 activation mechanisms, 17, 18 blood vessels/vascular structures, 362, 364 endothelial function, 379 endothelium, 385 smooth muscle, 365, 367, 368 GenBank data, 5 genomic structures, 11 growth factors and, 18 heteromeric channels, 32, 59, 60, 333 muscarinic receptor stimulated, 18

Index neuron pathﬁnding/migration, 4, 59 protein–protein interactions binding partner effects on channel localization and function, 335 IP3 receptor, 340 signal transduction, 17 translocation of, agonist-induced, 325–326 tyrosine phosphorylation, role in function of, 19, 20–21 TRPC7, 24, 59 blood vessels/vascular structures, 362, 364 heteromeric channels, 32, 333 protein–protein interactions binding partner effects on channel localization and function, 336 heteromeric channels, 333 signal transduction, 17 SOCE, 23, 34 tyrosine phosphorylation, role in function of, 19 trp and TRPL, 2, 15, 338, 364 TRPM (subfamily), 32, 234, 323, 363 blood vessels/vascular structures, 362 GenBank data, 6 genomic structures, 9, 12, 13 hetero and homomeric ion channels, 333 pore forming sequence similarity, 142 structure enzymatic domains, 355 MHRs, 355 pore domain, 352 TRP subfamilies, 125, 190 TRPM1, 5, 362 TRPM2 amino acid sequence alignment, 8 blood vessels/vascular structures, 362 calcium selectivity/nonselectivity, 4 GenBank data, 5 structure, 351, 355 TRPM3 blood vessels/vascular structures, 362 GenBank data, 5 osmotic sensitivity, 4 phylogenetic tree and regulatory inputs, 7 volume regulation, 142, 145–147 activation and pharmacological properties, 146–147 molecular features and tissue expression, 145–146 TRPM4 blood vessels/vascular structures, 362 endothelium, 369, 379, 380 smooth muscle, 368 GenBank data, 5 osmotic sensitivity, 4 phylogenetic tree and regulatory inputs, 7

463 pore domain, 352 voltage dependence, 353 TRPM4/TRPM5, 203–210 calcium selectivity/nonselectivity, 4 calcium sensitivity, 205–206 cloning, distribution, and structure, 204 function of, 208–209 ion permeability, unitary properties, and blockers, 204–205 models for physiological activation, 209 PIP2 regulation, 207–208 selectivity/nonselectivity of channels, 363 thermosensation, 272 chemical agonists, 280, 281 role of TRP channels, 277–278 vascular smooth muscle, 367 voltage activation, 206 TRPM5 blood vessels/vascular structures, 362 electrogenic properties, 18 GenBank data, 5 phylogenetic tree and regulatory inputs, 7 pore domain, 352 taste transduction channel, 4 TRPM6 blood vessels/vascular structures, 362 calcium selectivity/nonselectivity, 4 enzymatic domains, 355 GenBank data, 5 TRPM7 blood vessels/vascular structures, 362 calcium selectivity/nonselectivity, 4 GenBank data, 6 structure, 351, 355 TRPM8 (CMR1), 4, 177–186, 206, 208 blood vessels/vascular structures, 362 cloning of cold and menthol receptor, 180 and cold receptor properties, 180–182 cold sensing and menthol, 178 cooling compounds activating, 182–183 GenBank data, 6 icilin and, 171 mechanism of cold activation, 184–185 menthol receptor, hunt for, 178–180 phylogenetic tree and regulatory inputs, 7 regulation of currents, 183–184 role for, outside nervous system, 185 structure, pore domain, 352 temperature sensitivity, 288–289 allosteric model, 297–299 multistate nature of channel opening process, 295 thermodynamics, 289, 290, 296 voltage dependence, 291–292, 294

464

TRP Ion Channel Function in Sensory Transduction

thermosensation, 272 chemical agonists, 280, 281 role of TRP channels, 278–279 temperature sensors in mammals, 217 TRPV1 coexpression, 96 voltage dependence, 353 TRPML, 126, 234, 323, 363 discovery of, 3 genomic structures, 9, 11 TRP subfamilies, 190 TRPML1, 6, 8 TRPML2, 6 TRPML3, 164 TRPN (subfamily), 190, 323 ankyrin repeats, 353, 354 TRPA channels and, 152 TRPN1, 164 TRP-Nudix, 7 TRPP (subfamily), 190–191, 192, 234, 323 Caenorhabditis elegans mating and fertilization, 257–261 future directions, 261 intraﬂagellar transport of TRPV but not TRPP in cilia, 260 kinesin KLP-6 regulation, 259–260 PLAT binding partners, 260–261 polycystin ciliary localization and ADPKD, 257–259 discovery of, 3 GenBank data, 6 genomic structures, 9, 11, 13 hetero and homomeric ion channels, 333 nomenclature, 4 TRP subfamilies, 126 TRPP1, ciliated epithelia, 417 TRPP1 exons, 11 TRPP2 (polycystin-2), 144, 259 activation mechanisms and functional roles, 189–199 endoplasmic reticulum, 194 localization and trafﬁcking, 193 mechanosensitivity, 196–199 PKD1 complex activation, 195–196 PKD1 complex functions, 194–195 polycystic kidney disease, mutations causing, 191–193 signaling and regulation, 192 TRPP subfamily, 190–191, 192 amino acid sequence alignment, 8 ciliated epithelia, 417 KLP-6 colocalization, 259 and left-right asymmetry in mouse development, 260 TRPP3, 6, 190, 193 TRPP3a, 191

TRPP4, see PKD-REJ (receptor for egg jelly, TRPP4) TRPP5, 6, 190, 193 TRPV (subfamily), 32, 234, 323, 363 blood vessels/vascular structures, 362 Caenorhabditis elegans, 260; see also Caenorhabditis elegans ciliary localization, 259 discovery of, 3 Drosophila, 234–237; see also Drosophila, hearing and proprioception GenBank data, 6 genomic structures, 9, 10, 12, 13 hetero and homomeric ion channels, 333 mammalian serotonergic systems, 251 osmo- and mechanotransduction, channel functions in, 304 pore forming sequence similarity, 142 protease-activated receptors, 426 protease-activation in nervous system, 430–433 SOCE channels, 23 structure ankyrin repeats, 353, 354 pore domain, 352 TRP subfamilies, 126, 190 TRPV1 (VR1), 279 amino acid sequence alignment, 8 blood vessels/vascular structures, 362 calcium-dependent inactivation, 134 calmodulin binding sites, 133 coexpression with TRPA1, 279 discovery of, 3 GenBank data, 6 genomic structures, 12 mammalian serotonergic systems, 251 mechanotransduction, 222, 305–306 osmotransduction, 305–306 phylogenetic tree and regulatory inputs, 7 protease-activated receptor sensitization of TRPV channels, 430 structure ankyrin repeats, 354–355 oligomerization of cytosolic domains, 356 temperature sensitivity, 4, 288 allosteric model and, 299 multistate nature of channel opening process, 295 thermodynamics, 289, 290, 296 voltage dependence, 291–292, 294–295 thermosensation chemical agonists, 280, 281 role of TRP channels, 273–275 temperature sensors in mammals, 217–218 translocation of channels, 324, 327, 328 TRPA1 coexpression, 157

Index TRPM8 antagonists and, 183 TRPV1/4 heteromeric receptors, 309 voltage dependence, 353 TRPV1 receptors and signal transduction activators, 72–75 capsaicin and other agonists, 72–73 heat, noxious, 74–75 protons, 73–74 modulation by cellular components, 75–78 calmodulin and phosphorylation, 76–77 lipids, 77 PKA, phosphorylation by, 76 PKC, phosphorylation by, 76 reducing agent effects, 77–78 TRPV1 regulation, 86–98 activators, 89–90 coexpression on sensory neurons and overlapping activity, 96 diversity of behavior dissimilar structure-activity relationships, 91–92, 93 endogenous ligands, 92–94 endogenous ligands, 87–89 by mast cells, 91 by neuroendocrine factors, 95 NGF and inﬂammatory hyperalgesia, 90 pathological conditions, changes of expression under, 97 pharmacological overlap between TRPV1 and its relatives, 96–97 protein kinase modulation, 90–91 resting state, 89 splice variants, possible regulation by, 95 by steroids, serum, 95 subcellular compartments, shufﬂing between, 91 tissue and cellular expression of VR1 and its splice variants, 94 whole animal level, opposing actions at, 96 TRPV2 (VRL-1), 3, 251 blood vessels, smooth muscle, 362, 367–368 cardiovascular system, 389–394 discovery of, 3 GenBank data, 6 growth factor regulation, 18 IGF regulation, 105–112 calcium entry requirement for cell-cycle progression, 105–106 molecular identiﬁcation of channel, 107 property of channel, 106–107 regulatory process, 107–109 translocation of receptor, regulation of, 109–112 osmo- and mechanotransduction, channel functions in, 306–307

465 phylogenetic tree and regulatory inputs, 7 structure, ankyrin repeats, 354–355 temperature sensitivity, 4 thermosensation, 272, 288 chemical agonists, 280, 281 role of TRP channels, 275–276 temperature sensors in mammals, 217 TRPV3 (VRL-2), 97, 251 blood vessels/vascular structures, 362 GenBank data, 6 genomic structures, 12 osmo- and mechanotransduction, channel functions in, 307 phylogenetic tree and regulatory inputs, 7 temperature sensitivity, 4, 288 thermodynamics, 296 voltage dependence, 292 thermosensation, 272 chemical agonists, 280, 281 role of TRP channels, 276–277 temperature sensors in mammals, 217 voltage dependence, 353 TRPV4, 97, 192, 251 blood vessels/vascular structures, 362, 394 endothelium, 369, 370, 371, 377–386 smooth muscle, 365, 366 blood vessels/vascular structures, role in endothelial mechanotransduction, 377–386 antihypertensives, pharmacological potential, 386 calcium and endothelial function, 378–379 cardiovascular disease, 385–386 functional role, 383–385 nitric oxide dependence of vasodilatation in carotid arteries, 383 TRPs in endothelium, 379–380 TRPV4 in endothelium, 380–382 vasodilatation, 382–383 channel properties, 135–136 biophysical properties, 135–136 blockers, 136 ciliated epithelia, 413–418 GenBank data, 6 invertebrate counterparts, 198 kidney, 397–409 nociception, 4 osmo- and mechanotransduction, channel functions in evolutionary conservation of function, 303–304 tissue culture and animal data, 307–310 phylogenetic tree and regulatory inputs, 7 regulation of, 127–135

466

TRP Ion Channel Function in Sensory Transduction

calcium dependence, 133–134 hypotonic solutions and mechanical stimuli, mechanisms of, 130–131 interaction of different stimuli, 135 mechanosensitivity, 129–130 osmolarity, extracellular, 127–129 phorbol esters, activation by, 131–132 temperature, 132–133 signal transduction, 312 thermosensation/temperature sensitivity, 272, 288 chemical agonists, 280, 281 role of TRP channels, 276–277 temperature sensors in mammals, 217 voltage dependence, 294 tissue expression, 126–127 TRP subfamilies and nomenclature, 125–126 volume regulation, 142–145 activation and pharmacological properties, 143–144 and cellular function, 144–145 molecular features and tissue expression, 142–143 TRPV4 gating, molecular mechanisms, 113–121 channel activation mechanisms, 116–119 arachidonic acid and epoxygenase metabolites, 119 diacylglycerol and phorbol esters, 118 mechanosensitivity, 116–117 thermosensitivity, 117–118 emerging paradigms of gating, 120 molecular basis of selectivity, 119–120 structure, 114–116 amino terminus, 114, 115 carboxy terminus, 115, 116 membrane-spanning core and pore loop, 115–116 TRPV5 (CaT2, ECaC1) blood vessels/vascular structures, 362 GenBank data, 6 genomic structures, 12, 13 selectivity/nonselectivity of channels, 363 SOCE channels, 23 structure oligomerization of cytosolic domains, 356 pore domain, 352 translocation of channels, 327 TRPV6 (CaT1, ECaC2) blood vessels/vascular structures, 362 calcium-dependent inactivation, 134 calmodulin binding sites, 133 GenBank data, 6 genomic structures, 12, 13 selectivity/nonselectivity of channels, 363 SOCE channels, 23 structure

oligomerization of cytosolic domains, 356 pore domain, 352 translocation of channels, 327 Trypsin, 274, 424, 425 Trypsinogens, 425, 429 Tryptase, 91, 274, 424, 425, 426, 428 Tubulin, sensory cilia, 228 Type I sensory neurons, 217 Type II sensory neurons, 215, 216, 218 Tyrosine kinases neuron pathﬁnding/migration, 59 TRPC family role in function of, 19–24 voltage-gated calcium channel-independent calcium inﬂux, 19 TRPV1, 90–91 TRPV4, mechanisms of activation by hypotonic solutions and mechanical stimuli, 130

U U73122, 14 Uncoordinated (unc) mutants, 216, 231 Untranslated sequences, 9 Uridine triphosphate, 362, 365 Urinary bladder, 73, 94, 305 U73122, 59

V Vone1R family of vomeronasal receptors, 47 VAMP2, 326, 327, 328, 336, 339 Vanilloid binding, TRPV1, 70 Vanilloid channels, see TRPV1 (VR1) Vasculature, see Blood vessels Vasoactive intestinal polypeptide (VIP), 428 Vasoconstriction agonist-induced channel activation, 363–366 Vasodilation (vasodilatation), 379 cannabinoids and, 156 response to EETs mediated by channel activation, 366, 371 TRPV4 role, 382–383 antihypertensives, pharmacological potential, 386 cardiovascular disease, 385–386 nitric oxide dependence in carotid arteries, 383 Vasopressin receptors, 22 Vesicles, subcellular SNAREs, 320–321 TRPV2, 108, 111 Vestibular organs, TRPA1, 164, 165 Vinculin, 62

Index Vision photoreceptors, see Photoreceptor cells trichromatic, and pheromone signaling, 46, 47, 50 VOLT, TRPV4, 143 Voltage activation, TRPM4/TRPM5, 206 Voltage dependence structure and, 353 thermoTRP channel gating biophysics, 291–294 Voltage-dependent calcium channels, 7 neuron pathﬁnding/migration, 59, 60, 61 TRPV1 activation, 73 TRPV4, 118 vascular smooth muscle, 367 Voltage-dependent currents, TRPM8, 184–185 Voltage-dependent gating potassium channels, TRPC2 and, 48 thermoTRP channels, 291–294 TRPM4, PIP2 and, 208 TRPV1, 75, 275 Voltage-gated calcium channel conformational coupling, 15–16 TRPC channels and, 342 Volume regulation ciliated epithelia, 415–416 regulatory volume decrease (RVD), 307 signal transduction, 312 TRPM3, 145–147 activation and pharmacological properties, 146–147 molecular features and tissue expression, 145–146 TRPV4, 117, 142–145 activation and pharmacological properties, 143–144 and cellular function, 144–145 molecular features and tissue expression, 142–143

467 Vomeronasal organs, TRPC2 expression, 4, 32, 45–51 V1R family of vomeronasal receptors, 46 VR1, see TRPV1 (VR1) VRL-1, see TRPV2 (VRL-1) VRL-2, see TRPV3 (VRL-2) VROAC, see TRPV4 v-SNARE, see Soluble N-ethylmaleimidesensitive factor attachment protein receptors (SNAREs) VTTRL motif, 335 V2R family of vomeronasal receptors, 46, 48

W Wasabi, 222, 279, 289 Water balance, TRPV4 in systemic osmoregulation, 405–409 Water wire, 74 Whole animal level, TRPV1, 96

X Xenopus oocyte expression system, 182, 324–325 Xenopus spinal neurons, 4, 58 Xenopus TRP1 (xTRPC1), 58–59

Y yes-deﬁcient cells, 19, 21, 22 YF, 19

Z Zebraﬁsh, 152, 164 Zinc, 351 Zingerone, 73 Zona occludens 1 (ZO-1), 336, 337

COLOR FIGURE 1.6 Overview of some of the mechanisms that regulate or are thought to regulate the Gq-PLCβ triggered activation of TRPCs. A ligand of a Gq-coupled G-proteincoupled receptor (GPCR) is shown to activate the receptor’s Gq-activating function whereby Gq’s GDP is changed to GTP with concomitant dissociation of the heterotrimeric Gq protein into GTP-α plus the β-γ dimer. Both GTP-αq and β-γ independently activate β-type phospholipase Cs (PLCβs) leading to the hydrolysis of phosphatidylinositol bis-phosphate (PIP2) into inositol-trisphosphate (IP3) plus diacylglycerol (DAG). TRPC channels are depicted as being activated by three distinct mechanisms: (1) by DAG, presumably acting by direct interaction with the TRPC; (2) by the inositol trisphosphate receptor (IP3R), also thought to interact directly with the TRPC; and (3) possibly by STIM1, an ER Ca sensor that upon store depletion is translocated from the endoplasmic reticulum membrane (ER) to the plasma membrane (PM). The ﬁgure also highlights the negative regulation by PKC, activated by the cooperative interaction effect of DAG (generated by the action of the PLCβ) and Ca2+, originating ﬁrst from the store, and later from the extracellular milieu entering through the TRPC. CAMKs, Ca-calmodulin activated kinases; NOS, nitric oxide synthase; CN, calcineurin—also PP2B.

COLOR FIGURE 3.1 (A) The vomeronasal system: pheromone signals are detected by neurons of the vomeronasal organ (VNO) in the ventral nasal septum. VNO axons project to the accessory olfactory bulb (AOB), which in turn connects to nuclei of the vomeronasal amygdala in the limbic system. (B) Key signaling components of the vomeronasal organ: VNO neurons express one member of the two families of vomeronasal receptors V1Rs and V2Rs. V2Rs form a functional complex with the MHC class Ib molecules M10s. The activities of V1R- and V2R-expressing neurons require the expression of the TRPC2 channel. (C) Localization of rTRPC2 to the sensory microvilli of VNO neurons: anti-TRPC2 immunoﬂuorescence on sections of rat VNO demonstrates the localization of TRPC2 to the luminal surface of the VNO neurepithelium where pheromone-induced signaling is likely to occur. (Modiﬁed from reference 20.) (D) TRPC2 in primate evolution. Mutations that disrupt the opening of reading frame (ORF) of the TRPC2 protein were mapped to the time in primate evolution at which they ﬁrst occurred. The ﬁrst mutations (6 and 9) occurred in the ancestors of Old World monkeys and apes, at the same time that these animals developed trichromatic vision. Howler monkeys have independently evolved trichromatic vision but have an intact TRPC2 gene, indicating the trichromatic vision is not in itself sufﬁcient to replace pheromone signaling.37 (Modiﬁed from reference 21.) (E) The role of VNO signaling in gender discrimination and aggression: behavioral analysis of TRPC2-/- males reveals the essential role of the VNO in controlling the sex speciﬁcity of reproductive behavior, and in male-male aggression. Other sensory cues, most likely olfactory, are essential to trigger mating behavior in mice.

COLOR FIGURE 5.1 (A) A schematic diagram of a TRPV1 subunit in a bilayer. The subunit has six transmembrane domains (red) and a pore loop. The functional TRPV1 receptor is believed to form a tetramer. Residues involved in vanilloid binding are shown in orange. “A” indicates ankyrin repeats shown in yellow. Residues susceptible of phosphorylation are shown in green. Two calmodulin-binding regions in the N- and C-termini are indicated by “CaM.” Blue residues in the “P-loop” represent protonatable amino acids. Cysteine residues in the Ploop are susceptible of reduction and are indicated by the color purple. PIP2 is shown to bind to the region indicated in the C-terminus. The TRP box represents the TRP domain. (Modiﬁed from Ferrer-Montiel et al., 2004 and from Tominaga and Tominaga, 2005.) (B) Role of metabolic pathways of anandamide and arachidonic acid in TRPV1 activation. The fatty acid amide hydrolase (FAAH) hydrolyzes anandamide (AEA) to produce arachidonic acid (AA) and ethanolamide. AA is oxygenated by lipoxygenase enzymes to produce TRPV1 agonists, 12- and 15-HPETE, 5-HETE and leukotriene B4 (LTB4) (Hwang et al., 2000). AEA, a substrate for lipoxygenase, yields equivalent HPETE ethanolamides (HPETEE) and HETE ethanolamides (HETEE) that are proposed to be TRPV1 agonists (Craib et al., 2001). The lipoxygenase products of anandamide act as potent inhibitors of FAAH (Maccarrone et al., 2000). PKC activates the TRPV1 receptor directly (Premkumar and Ahern, 2000; Olah et al., 2002), but it also sensitizes the receptor to other agonists (Vellani et al., 2001). AEA directly activates PKC (De Petrocellis et al., 1995; Premkumar and Ahern, 2000). CB1 receptor activation is coupled to PLC stimulation, PIP2 hydrolysis and release of the TRPV1 receptor from the inhibitory effect of this compound (see Hermann et al., 2003). AEA production occurs through phosphodiesterase-mediated cleavage of a phospholipid precursor, NAPE (Narachidonoyl-phosphatidylethanolamine), in calcium-dependent fashion (DiMarzo et al., 1994). AEA is synthesized in response to TRPV1 receptor activation in cultured neurons (Ahluwalia et al., 2003). (Modiﬁed from Ross, 2003.)

COLOR FIGURE 14.1 Signaling and regulation of TRPP2 channels. This illustration depicts the different models for the localization-dependent functions of TRPP2. TRPP2 mediates Ca2+ inﬂux at the plasma membrane (PM) and the ciliary membrane (cilium), where it functions in a heteromultimeric protein complex with PKD1 (1, 3) and possibly with other members of the TRP channel superfamily (TRPC1, TRPV4) (2). PKD1 is known to activate multiple signaling pathways via G proteins, a process that can be regulated by physical interaction with TRPP2. Calcium inﬂux induced by shear ﬂow in renal epithelial cells is triggered by bending the primary cilium and seemingly requires the PKD1–TRPP2 complex in the luminal cilium (3). Mechanical stress activates PKD1–TRPP2 complexes to allow Ca2+ inﬂux either in the shaft or in the base of the primary cilium. TRPP2 acts as a Ca2+-release channel in the endoplasmic reticulum (ER), where it might interact with and regulate IP3Rs (4). Serinephosphorylated TRPP2 sequesters Id2 in the cytoplasm and normally prevents Id2 from entering the nucleus and binding to E-proteins (5). PKD1 can undergo a proteolytic cleavage that releases its C-terminal tail, which translocates to the nucleus and activates the transcription factor AP-1 (6). Note that we hypothetized that PKD1 targeted to the primary cilium is also cleaved at its GPS site. See text for more details. Abbreviations: PKC, protein kinase C; PKA, cAMP-dependent protein kinase; NFAT, nuclear factor of activated T-cells; PI3K, phosphatidyl-inositol 3-kinase; MEK, mitogen-activated protein kinase/ERK kinase; ERK, extracellular signal-regulated kinase.

A

Osmolarity and Gentle touch

OSM-9 GPCR

OCR-2

Lipid

G

B

COLOR FIGURE 18.1 Polymodality and speciﬁcity of OCR-2/OSM-9 in vivo. (A) OCR2/OSM-9 in different sensory functions are regulated by distinct mechanisms. OCR-2 and OSM-9 likely form a heteromeric channel and are located in the sensory cilia and plasma membrane of four classes of chemosensory neurons in the sensory organ amphids. The membrane region is shown in gray. The transmembrane domains of OCR-2 and OSM-9 are shown as oval columns, the rectangular bar represents the ankyrin motifs, and the ball structure represents the N-terminal region preceding the ankyrin motifs. OCR-2/OSM-9 function in diacetyl sensation depends on the G-protein-coupled receptor ODR-10, the G protein ODR3, and the signals from polyunsaturated fatty acids, but is not affected by the OCR-2(G36E) mutation in the ball. OCR-2/OSM-9 sensation to external osmolarity and gentle touch at the nose requires the G protein ODR-3, the signals from polyunsaturated fatty acids, and the determinants located in the ball region of OCR-2. Upregulation of tph-1 expression in the ADF neurons is governed by the determinants located in the ball of OCR-2. Abbreviations: G, heterotrimeric G protein; GPCR, G-protein-coupled receptor. (B) Photomicrographs of the serotonergic chemosensory neurons ADF expressing OCR-2. Both the wild-type OCR-2 and the mutant OCR-2(G36E) proteins are tagged to the FLAG epitope, and the transgenic worms were stained with anti-FLAG antibody. Notice that both wild-type OCR-2 and OCR-2(G36E) are expressed in the cilia (arrowheads) and the cell bodies.

COLOR FIGURE 20.1 (A) Temperature ranges activating thermosensitive TRP channels. (B) Phylogenetic relationship among the mammalian TRPV, TRPM, and TRPA channels with two Drosophila TRPA channels. Red, orange, and blue squares indicate channels activated by high heat, warm stimuli, and cold stimuli, respectively.

COLOR FIGURE 23.3 TRPV1 is translocated to the cell surface by regulated exocytosis. (A) TRPV1 colocalizes in neuronal vesicles with the v-SNARE protein VAMP2, as evidenced from the immunocytochemistry.39 (B) BoNTA abrogates the PKC-induced translocation of TRPV1 to the plasma membrane, as concluded from biotinylation of surface-expressed receptors.39,40 TPA denotes 12-O-tetradecanoylphorbol-13-acetate, a potent agonist of PKC.

COLOR FIGURE 25.1 Structures of TRP channel-building blocks. The transmembrane domain of the Shaker channel (2A798), viewed from the intracellular side of the membrane (A) and from its side (B). The four subunits have distinct colors, and magenta spheres represent potassium ions in the selectivity ﬁlter. The S1–S4 helices form the sensor domain, connected to the S5–S6 pore domain through the S4–S5 linker. The S4–S5 linker forms a ring around the C-terminal end of the S6 helices and may trigger motion of the S6 helices to open or close the pore in response to voltage changes. Each sensor domain abuts the pore domain of the neighboring subunit, with their respective S4 and S5 helices in close contact. (C) Structure of the six ankyrin repeats in the N-terminal region of TRPV2 (2ETA44). Many TRP channels have ankyrin repeats in their N-terminal cytosolic regions. (D) Structure of the TRPM7 α-kinase domain, located at the C-terminal end of the channel (1IA951). An AMPPNP molecule is located in the active site, and a structural zinc ion is displayed as a grey sphere. (E) Structure of human NUDT9 with a ribose-5-phosphate molecule and two magnesium ions in the active site (1QVJ53). The C-terminal end of TRPM2 has 39 percent similarity to NUDT9.

COLOR FIGURE 30.2 Confocal microscopy localization of TRPV4 to motile cilia in a tissue section of the female hamster oviduct. (Left) Nomarsky image. (Right) Detection of TRPV4 (green) and TOPRO-3 nuclear staining (blue). The antibody used to detect TRPV4 is described in references 6, 8, and 22.

Mast Cells Epithelium

Circulation

Neurons

INJURY AND INFLAMMATION FVIIa, FXa

Tryptase Trypsin IV Trypsin IV SLIGRL 2 1

PAIN TRANSMMISION TRPV Channels

PAR

7 2 3

CLR/RAMP1

CGRP

Sensory nerve ending

SP

CLR/RAMP1 SP CGRP 6

NK1R Spinal neuron

NK1R 5 4 Arteriolar dilation

Venular permeability Granulocyte infiltration

NEUROGENIC INFLAMMATION

COLOR FIGURE 31.1 Protease cleavage and activation of PAR2 on primary spinal afferent neurons to cause neurogenic inﬂammation and pain. (1) Injury and inﬂammation trigger the generation and release of proteases from mast cells (tryptase), epithelial cells, and neurons (trypsins), and the circulation (FVIIa, FXa) that can cleave PAR2 at the peripheral projections of primary spinal efferent neurons. (2) Cleavage exposes the tethered ligand domain (SLIGRL in mice and rats), which binds conserved regions of the extracellular loop II to activate the receptor. (3) PAR2 activation increases [Ca2+]i and releases CGRP and SP in peripheral tissues. (4) CGRP interacts with the type 1 CGRP receptor (a heterodimer of calcitonin receptor-like receptor, CLR, and receptor activity modifying protein 1, RAMP1) on arterioles to induce dilation and hyperemia. (5) SP interact with the neurokinin 1 receptor (NK1R) on endothelial cells of postcapillary venules to induce gap formation and plasma extravasation. Together, these induce neurogenic inﬂammation. (6) The same stimuli release SP and CGRP in the spinal cord dorsal horn to induce pain transmission. (7) PAR2 sensitize TRPV1 and TRPV4 channels, which causes hyperalgesia to thermal and mechanical stimuli, respectively.