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


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


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 -


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


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


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


V S

A A T

K K S

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


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 - - - -


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.


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).



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


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


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


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.


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


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.


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


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.


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


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


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


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


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.

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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

Multiple Mechanisms of TRPC Activation

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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


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


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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.

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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.


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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.

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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


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.

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(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


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


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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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

58


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


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


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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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.

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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

TRPV1 Receptors and Signal Transduction

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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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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.


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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).

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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


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.

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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 (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.

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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

TRPV2: A Calcium-Permeable Cation Channel

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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.

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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

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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 (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

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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

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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.

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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

OF

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.


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.

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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.

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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

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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

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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

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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.

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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.

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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 (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 (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
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


383

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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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.

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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).

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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

392


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


393

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

394


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.


395

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.

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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


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

The Role of TRPV4 in the Kidney

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,

400


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


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


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.


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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


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


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


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

408


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


409

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.

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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.

30

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

The TRPV4 Channel in Ciliated Epithelia

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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


417

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.

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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

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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

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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.

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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


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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.

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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


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

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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


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.

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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


431

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

432


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


433

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.

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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


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


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


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


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


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


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


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


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


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


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 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


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


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.

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

Transduction Channels in Sensory Cells

Methods in Genomic Neuroscience (Frontiers in Neuroscience)

Methods in Insect Sensory Neuroscience

In Vivo Optical Imaging of Brain Function (Frontiers in Neuroscience)

In Vivo Optical Imaging of Brain Function (Frontiers in Neuroscience)

Molecular and Cellular Signaling

Molecular and Cellular Signaling

Molecular and Cellular Signaling

Molecular and Cellular Insights to Ion Channel Biology

Cellular and Molecular Methods in Neuroscience Research

Cellular and Molecular Methods in Neuroscience Research

Cellular and Molecular Methods in Neuroscience Research

Biomedical Imaging in Experimental Neuroscience (Frontiers in Neuroscience)

Advances in the Neuroscience of Addiction (Frontiers in Neuroscience)

Biomedical Imaging in Experimental Neuroscience (Frontiers in Neuroscience)

Serotonin Receptors in Neurobiology (Frontiers in Neuroscience)

Advances in the Neuroscience of Addiction (Frontiers in Neuroscience)

Methods of Behavior Analysis in Neuroscience (Frontiers in Neuroscience)

Methods in Drug Abuse Research: Cellular and Circuit Level Analyses (Methods and New Frontiers in Neuroscience)

Ion Channel Localization (Methods in Pharmacology and Toxicology)

Hydrogen-ion wave function

Photomorphogenesis in Plants and Bacteria: Function and Signal Transduction Mechanisms

Ion Channel Regulation (Advances in Second Messenger and Phosphoprotein Research)

Cellular Nanoscale Sensory Wave Computing

Lipid-Mediated Signaling (Methods in Signal Transduction Series)

The Neurobiology of Olfaction (Frontiers in Neuroscience)

Cellular Implications of Redox Signaling

The Neurobiology of Olfaction (Frontiers in Neuroscience)

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

Transduction Channels in Sensory Cells

Methods in Genomic Neuroscience (Frontiers in Neuroscience)

Methods in Insect Sensory Neuroscience

In Vivo Optical Imaging of Brain Function (Frontiers in Neuroscience)

In Vivo Optical Imaging of Brain Function (Frontiers in Neuroscience)

Molecular and Cellular Signaling

Molecular and Cellular Signaling

Molecular and Cellular Signaling

Molecular and Cellular Insights to Ion Channel Biology

Cellular and Molecular Methods in Neuroscience Research

Cellular and Molecular Methods in Neuroscience Research

Cellular and Molecular Methods in Neuroscience Research

Biomedical Imaging in Experimental Neuroscience (Frontiers in Neuroscience)

Advances in the Neuroscience of Addiction (Frontiers in Neuroscience)

Biomedical Imaging in Experimental Neuroscience (Frontiers in Neuroscience)

Serotonin Receptors in Neurobiology (Frontiers in Neuroscience)

Advances in the Neuroscience of Addiction (Frontiers in Neuroscience)

Methods of Behavior Analysis in Neuroscience (Frontiers in Neuroscience)

Methods in Drug Abuse Research: Cellular and Circuit Level Analyses (Methods and New Frontiers in Neuroscience)

Ion Channel Localization (Methods in Pharmacology and Toxicology)

Hydrogen-ion wave function

Photomorphogenesis in Plants and Bacteria: Function and Signal Transduction Mechanisms

Ion Channel Regulation (Advances in Second Messenger and Phosphoprotein Research)

Cellular Nanoscale Sensory Wave Computing

Lipid-Mediated Signaling (Methods in Signal Transduction Series)

The Neurobiology of Olfaction (Frontiers in Neuroscience)

Cellular Implications of Redox Signaling

The Neurobiology of Olfaction (Frontiers in Neuroscience)

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