Structure, Function and Modulation of Neuronal Voltage-Gated Ion Channels

STRUCTURE, FUNCTION, AND MODULATION OF NEURONAL VOLTAGEGATED ION CHANNELS

STRUCTURE, FUNCTION, AND MODULATION OF NEURONAL VOLTAGEGATED ION CHANNELS VALENTIN K. GRIBKOFF, PhD Knopp Neurosciences Inc., and The Department of Pharmacology, Yale University School of Medicine

LEONARD K. KACZMAREK, PhD The Department of Pharmacology Yale University School of Medicine

Copyright Ó 2009 by John Wiley & Sons, Inc. All rights reserved Published by John Wiley & Sons, Inc., Hoboken, New Jersey Published simultaneously in Canada No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, scanning, or otherwise, except as permitted under Section 107 or 108 of the 1976 United States Copyright Act, without either the prior written permission of the Publisher, or authorization through payment of the appropriate per-copy fee to the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, (978) 750-8400, fax (978) 750-4470, or on the web at www.copyright.com. Requests to the Publisher for permission should be addressed to the Permissions Department, John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, (201) 748-6011, fax (201) 748-6008, or online at http://www.wiley.com/go/permission. Limit of Liability/Disclaimer of Warranty: While the publisher and author have used their best efforts in preparing this book, they make no representations or warranties with respect to the accuracy or completeness of the contents of this book and specifically disclaim any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives or written sales materials. The advice and strategies contained herein may not be suitable for your situation. You should consult with a professional where appropriate. Neither the publisher nor author shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. For general information on our other products and services or for technical support, please contact our Customer Care Department within the United States at (800) 762-2974, outside the United States at (317) 572-3993 or fax (317) 572-4002. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic formats. For more information about Wiley products, visit our web site at www. wiley.com. Library of Congress Cataloging-in-Publication Data: Gribkoff, Valentin K. Structure, function, and modulation of neuronal voltage-gated ion channels / Valentin K. Gribkoff, Leonard K. Kaczmarek. p. cm. Includes index. ISBN 978-0-470-42989-1 (cloth) 1. Ion channels. 2. Ion channels–Effect of drugs on. 3. Drug development. 4. Molecular pharmacology. I. Kaczmarek, Leonard K. II. Title. QH603.I54.G75 2009 6150 .10724–dc22 2008046707 ISBN: 9780471930136

Printed in the United States of America 10 9 8 7 6 5 4 3 2 1

CONTENTS

Preface

ix

Contributors

xi

PART I 1

NEURONAL VOLTAGE-GATED ION CHANNEL FUNCTIONS

1

Neuronal L-Type Voltage-Gated Calcium Channels

3

Alexander Scriabine and David J. Triggle

2

Voltage-Gated N-Type and T-Type Calcium Channels and Excitability Disorders

35

Elizabeth Tringham and Terrance P. Snutch

3

Voltage-Gated Sodium Channels: Multiple Roles in the Pathophysiology of Pain

67

Sulayman D. Dib-Hajj, Bryan C. Hains, Joel A. Black, and Stephen G. Waxman

4

The Role of Ion Channels in the Etiology and Development of Gliomas

105

Amy K. Weaver and Harald Sontheimer

5

Shaker Family Kv1 Voltage-Gated Potassium Channels in Mammalian Brain Neurons

127

Helene Vacher and James S. Trimmer

v

vi

6

CONTENTS

Unique Mitochondrial Ion Channels: Roles in Synaptic Transmission and Programmed Cell Death

155

Elizabeth A. Jonas

7

Regulation of Neuronal Excitability by the Sodium-Activated Potassium Channels Slick (SLO2.1) and Slack (SLO2.2)

193

Valentin K. Gribkoff and Leonard K. Kaczmarek

PART II MODULATORY MECHANISMS AND INFLUENCES ON NEURONAL VOLTAGE-GATED ION CHANNEL FUNCTION 8

Alternative Splicing of Neuronal Cav2 Calcium Channels

217 219

Diane Lipscombe, Summer E. Allen, Annette C. Gray, Spiro Marangoudakis, and Jesica Raingo

9

Effect of Hypoxia/Ischemia on Voltage-Dependent Channels

251

Xiang Q. Gu, Hang Yao, and Gabriel G. Haddad

10

In Vivo Roles of Ion Channel Regulatory Protein Complexes in Neuronal Physiology and Behavior

279

Smitha Reddy, Mohammad Shahidullah, and Irwin B. Levitan

11

Regulation of Neuronal Ion Channels by G-Protein-Coupled Receptors in Sympathetic Neurons

291

Mark S. Shapiro and Nikita Gamper

12

BK Channels: Regulation of Expression and Physiological Impact

317

Pallob Kundu, Abderrahmane Alioua, Yogesh Kumar, Rong Lu, Jimmy W. Ou, Enrique Sanchez-Pastor, Min Li, Enrico Stefani, and Ligia Toro

13

Structural Basis for Auxiliary KChIP Modulation of Kv4 Channels

343

KeWei Wang and Jijie Chai

PART III 14

DRUG DISCOVERY TARGETS AND TECHNOLOGY

Sodium Channel Blockers for the Treatment of Chronic Pain

363 365

Mark R. Bowlby and Edward Kaftan

15

Neuronal Kv7 Potassium Channels as Emerging Targets for the Treatment of Pain Steven I. Dworetzky and Valentin K. Gribkoff

393

CONTENTS

16

Small-Molecule Modulators of Large-Conductance, Calcium-Activated (BK) Channels

vii

423

John E. Starrett Jr.

17

High-Throughput Screening Technologies in Ion Channel Drug Discovery

443

Edward B. Stevens, Andrew D. Whyment, and J. Mark Treherne

Index

467

PREFACE

This book began as an idea in 2000 at a Keystone Conference on potassium channels that was organized by us, and the idea was subsequently expanded to include contributions on neuronal voltage-gated ion channels of all types. In 2005, after hosting a symposium on ion channels as drug discovery targets, one of us (V.G.) was invited by the editors at John Wiley & Sons to organize a book primarily focusing on ion channels as drug targets. We later decided that we would follow through on our original plan to publish a book that looked at neuronal voltage-gated ion channels from multiple perspectives, including the translational perspective of drug discovery and development. This was an ambitious project, and from the outset our plan was not to cover every facet of research in those areas encompassed by the title of this book, but to include chapters that were in-depth reviews of selected topics relevant to the structure, function, and modulation of these important ion channel families. We hoped that while the individual chapters would prove interesting and useful to established ion channel scientists, our priority was to provide students of this important and exciting field glimpses into a number of interesting areas worthy of their further study. What we have arrived at after more than 2 years of organizing, inviting, cajoling (gently), writing, and editing are 17 chapters from leading authorities in their respective fields, either in academia or industry. Each of these is either a comprehensive review of a particular voltage-gated ion channel member, such as the review by Scriabine and Triggle on “Neuronal L-type voltage-gated calcium channels” (Chapter 1), or is an in-depth review of a topic relevant to multiple voltage-gated ion channel subfamilies, such as the chapter by Shapiro and Gamper on the “Regulation of ion channels by G-proteincoupled receptors” (Chapter 11). The book is divided into three general sections: (1) Neuronal voltage-gated ion channel functions, (2) Modulatory mechanisms and

ix

x

PREFACE

influences on voltage-gated ion channel activity, and (3) Drug discovery targets and technology. There is some overlap in both chapter content and section assignment, but these divisions should serve to orient the reader to groups of chapters of similar scope. We trust that the reader will find these contributions interesting, useful for future reference and helpful in stimulating further discussion and research.

VALENTIN K. GRIBKOFF, PhD Knopp Neurosciences Inc., and The Department of Pharmacology, Yale University School of Medicine

LEONARD K. KACZMAREK, PhD The Department of Pharmacology, Yale University School of Medicine

CONTRIBUTORS

Abderrahmane Alioua, UCLA, Department of Anesthesiology, Division of Molecular Medicine, USA Summer E. Allen, Department of Neuroscience, Brown University, Providence, RI 02912, USA Joel A. Black, Department of Neurology and Center for Neuroscience and Regeneration Research, Yale University School of Medicine, New Haven, CT 06510, USA; Rehabilitation Research Center, VA Connecticut Healthcare System, West Haven, CT 06516, USA Mark R. Bowlby, Discovery Neuroscience, Wyeth Research, Princeton, NJ 08543, USA Jijie Chai, National Institute of Biological Sciences, No. 7 Science Park Road, Beijing 102206, China Sulayman D. Dib-Hajj, Department of Neurology and Center for Neuroscience and Regeneration Research, Yale University School of Medicine, New Haven, CT 06510, USA; Rehabilitation Research Center, VA Connecticut Healthcare System, West Haven, CT 06516, USA Steven I. Dworetzky, Knopp Neurosciences Inc., 2100 Wharton Street, Suite 615, Pittsburgh, PA 15203, USA Nikita Gamper, Institute of Membrane and Systems Biology, Faculty of Biological Sciences, The University of Leeds, Leeds LS2 9JT, UK

xi

xii

CONTRIBUTORS

Annette C. Gray, Department of Biology, Brandeis University, Waltham, MA 02454, USA Valentin K. Gribkoff, Department of Pharmacology, Yale University School of Medicine, New Haven, CT 06520, USA; Discovery Research, Knopp Neurosciences Inc., 2100 Wharton Street, Suite 615, Pittsburgh, PA 15203, USA Xiang Q. Gu, Department of Pediatrics, 9500 Gilman Drive, University of California at San Diego, San Diego, CA 92093-0735, USA Gabriel G. Haddad, Department of Pediatrics, 9500 Gilman Drive, University of California at San Diego, San Diego, CA 92093-0735, USA; Department of Neuroscience, 9500 Gilman Drive, University of California at San Diego, San Diego, CA 92093-0735, USA; The Rady Children’s Hospital-San Diego, 3020 Children’s Way, San Diego, CA 92123, USA Bryan C. Hains, Department of Neurology and Center for Neuroscience and Regeneration Research, Yale University School of Medicine, New Haven, CT 06510, USA; Rehabilitation Research Center, VA Connecticut Healthcare System, West Haven, CT 06516, USA Elizabeth A. Jonas, Department of Internal Medicine, Yale University, New Haven, CT 06520, USA Leonard K. Kaczmarek, Department of Pharmacology, Yale University School of Medicine, New Haven, CT 06520, USA Edward Kaftan, Discovery Neuroscience, Wyeth Research, Princeton, NJ 08543, USA Yogesh Kumar, UCLA, Department of Anesthesiology, Division of Molecular Medicine, USA Pallob Kundu, UCLA, Department of Anesthesiology, Division of Molecular Medicine, USA Irwin B. Levitan, Department of Neuroscience, University of Pennsylvania School of Medicine, 215 Stemmler Hall, 3450 Hamilton Walk, Philadelphia, PA 191046074, USA Min Li, UCLA, Department of Anesthesiology, Division of Molecular Medicine, USA Diane Lipscombe, Department of Neuroscience, Brown University, Providence, RI 02912, USA Rong Lu, UCLA, Department of Anesthesiology, Division of Molecular Medicine, USA Spiro Marangoudakis, Department of Neuroscience, Brown University, Providence, RI 02912, USA

CONTRIBUTORS

xiii

Jimmy W. Ou, UCLA, Department of Anesthesiology, Division of Molecular Medicine, USA Jesica Raingo, Department of Neuroscience, Brown University, Providence, RI 02912, USA Smitha Reddy, Department of Neuroscience, University of Pennsylvania School of Medicine, 215 Stemmler Hall, 3450 Hamilton Walk, Philadelphia, PA 191046074, USA Enrique Sanchez-Pastor, UCLA, Department of Anesthesiology, Division of Molecular Medicine, USA Alexander Scriabine, Department of Pharmacology, Yale University School of Medicine, New Haven, CT 06520, USA Mohammad Shahidullah, Department of Neuroscience, University of Pennsylvania School of Medicine, 215 Stemmler Hall, 3450 Hamilton Walk, Philadelphia, PA 19104-6074, USA Mark S. Shapiro, Department of Physiology, University of Texas Health Science Center at San Antonio, MS7756, San Antonio, TX 78229, USA Terrance P. Snutch, Neuromed Pharmaceuticals, Rm 301, 2389 Health Sciences Mall, Vancouver, BC, Canada; Michael Smith Laboratories, University of British Columbia, Vancouver, BC, Canada Harald Sontheimer, Department of Neurobiology and Center for Glial Biology in Medicine, The University of Alabama, Birmingham, AL 35294, USA John E. Starrett Jr., Discovery Chemistry, Bristol-Myers Squibb Co., 5 Research Parkway, Wallingford, CT 06492, USA Enrico Stefani, UCLA, Department of Anesthesiology, Division of Molecular Medicine, USA; UCLA, Department of Physiology, USA; UCLA, Department of Molecular and Medical Pharmacology, USA; UCLA, Cardiovascular Research Laboratory, USA Edward B. Stevens, Pfizer Global Research and Development, Sandwich Laboratories (IPC 351), Sandwich, Kent, CT13 9NJ, UK David J. Triggle, School of Pharmacy and Pharmaceutical Sciences, State University of New York, Buffalo, NY 14260, USA James S. Trimmer, Department of Neurobiology, Physiology and Behavior, College of Biological Sciences, University of California, Davis, CA 95616, USA; Department of Physiology and Membrane Biology, School of Medicine, University of California, Davis, CA 95616, USA Elizabeth Tringham, Neuromed Pharmaceuticals, Rm 301, 2389 Health Sciences Mall, Vancouver, BC, Canada

xiv

CONTRIBUTORS

Ligia Toro, UCLA, Department of Anesthesiology, Division of Molecular Medicine, USA; UCLA, Department of Molecular and Medical Pharmacology, USA; UCLA, Cardiovascular Research Laboratory, USA; UCLA, Brain Research Institute, USA J. Mark Treherne, Pfizer Global Research and Development, Sandwich Laboratories (IPC 351), Sandwich, Kent, CT13 9NJ, UK Stephen G. Waxman, Department of Neurology and Center for Neuroscience and Regeneration Research, Yale University School of Medicine, New Haven, CT 06510, USA; Rehabilitation Research Center, VA Connecticut Healthcare System, West Haven, CT 06516, USA Amy K. Weaver, Department of Neurobiology and Center for Glial Biology in Medicine, The University of Alabama, Birmingham, AL 35294, USA Helene Vacher, Department of Neurobiology, Physiology and Behavior, College of Biological Sciences, University of California, Davis, CA 95616, USA KeWei Wang, Neuroscience Research Institute and Department of Neurobiology, Key Laboratory for Neuroscience of the Ministry of Education, Center for Protein Sciences, Peking University Health Science Center, 38 Xueyuan Road, Beijing 100083, China Andrew D. Whyment, Pfizer Global Research and Development, Sandwich Laboratories (IPC 351), Sandwich, Kent, CT13 9NJ, UK Hang Yao, Department of Neuroscience, 9500 Gilman Drive, University of California at San Diego, San Diego, CA 92093-0735, USA

FIGURE 4.6 (a) Model representing glioma cell shape and volume-adaptive changes that occur during invasion in spatially restricted conditions. These changes, accompanied by water loss and cytoskeletal rearrangements, are mediated by ion fluxes through Cl
and Kþ ion channels and other ion transport mechanisms. (See text for full caption.)

FIGURE 4.7 Axial view of T1-Wc, (a) coregistered (b) and SPECT (c) (day 8) images. Longterm retention of 131I-TM-601 radioactivity at tumor site was observed in all patient scans (Hockaday et al., 2005).

FIGURE 5.1 Immunofluorescence staining of Kv1a and Kvb subunits in mammalian brain. (a) Staining for Kv1.1 (blue), Kv1.2 (green), and Kvb2 (red) in adult rat cerebellar cortex. Note overlap of all three fluors yielding white signal in basket cell terminals (arrowheads). (b) Staining for Kv1.2 (blue) in adult rat optic nerve (green ¼ Nav channels, red ¼ Caspr). Image courtesy of Dr. Matthew Rasband. (c) Staining for Kv1.4 (green) in the axon initial segment of a cultured hippocampal neuron (blue ¼ MAP2). Image courtesy of Drs. Yasuhiro Ogawa and Matthew Rasband. (d) Kv1.2 (red) in adult rat hippocampus (green ¼ Kv2.1). Arrowheads point to prominent Kv1.2 staining in perforant path presynaptic terminals in the middle molecular layer of the dentate gyrus.

FIGURE 5.2 Structure of Kv1a and Kvb subunits. (a) Cartoon of the predicted topology of Kv1a subunits. The predicted transmembrane topology of a Kv1a subunit and sites defined as important determinants of intracellular trafficking and polarized expression are shown schematically. T1, tetramerization domain important for Kvb-mediated intracellular and polarized trafficking of Kv1a subunits; NLG, N-linked glycosylation site of Kv1.1–Kv1.5; ERR, ER retention motif found in Kv1.1, Kv1.2, and Kv1.6; FTS, forward trafficking signal in Kv1.4; PDZ BD, PDZ-binding motif in all Kv1a subunits; Y132, tyrosine residue critical for tyrosine kinasedependent suppression of Kv1.2; S440/S441, serine residues critical for phosphorylationdependent trafficking of Kv1.2. (b) Crystal structure of the Kv1.2/Kvb2 complex (Protein Data Bank accession number 2A79). (See text for full caption.)

FIGURE 5.3 Subcellular localization of homotetrameric Kv1a subunits in COS-1 cells. Recombinant Kv1 expression monkey kidney fibroblast COS-1 cells. Intact cells were stained with ectodomain-directed antibodies (red), and then detergent permeabilized and stained with cytoplasmic domain-directed antibodies (green). (a) Kv1.1, (b) Kv1.4, (c and d) Kv1.2.

FIGURE 8.2 A large number of proteins and molecules act in concert to regulate alternative pre-mRNA splicing. In this hypothetical model of mutually exclusive splicing, all neurons express a number of regulatory proteins that either repress or enhance inclusion of one of the exon pair. (See text for full caption.)

FIGURE 8.3 Sites of alternatively spliced exons mapped on CaV2.1 and CaV2.2 proteins and the CaV2.2 gene. (a) and (b) illustrate the tetrameric structure and the twodimensional transmembrane topology of CaV2.1 and CaV2.2 proteins, respectively. Approximate locations of alternatively spliced exons are highlighted with blue and green triangles and are numbered. The four main structural domains (I, II, III, and IV) are labeled and the constitutively expressed exons encoding each domain shown with black lines. (c) illustrates the human CaV2.2 gene. Exons are denoted as solid boxes and introns as lines. (See text for full caption.)

FIGURE 8.4 Proteins that interact with the Cav2.2 subunit mapped onto a two-dimensional model of the channel. Several proteins interact with Cav2.2 channels, most notably in the II–III intracellular linker and C-terminus. A number of proteins associated with the synaptic machinery, and with synaptic transmission in general, interact with the II–III linker of Cav2.2, including CSP (cysteine string protein) (Miller et al., 2003), SNAP-25 (Catterall, 1999), synaptotagmin 1 (Sheng et al., 1997), syntaxin 1A (Sheng et al., 1994), huntingtin (Swayne et al., 2005), RGS12 (Richman and Diverse-Pierluissi, 2004), and b-arrestin (Puckerin et al., 2006). The asterisk (*) denotes proteins whose binding is either known or expected to be modified by alternative splicing. (See text for full caption.)

CaV2.2e[37b]

GPCR

CaV2.2e[37a]

GPCR

AP

ICa

FIGURE 8.5 Hypothesized physiological impact of alternative pre-mRNA splicing at the e37a/e37b site in CaV2.2. (See text for full caption.)

FIGURE 11.1 M-current in sympathetic neurons. (a) Biophysical properties of heterologously expressed M-channels. Kv7.2/7.3 heteromultimers were expressed in CHO cells and recorded from under whole-cell voltage-clamp. Currents were evoked by a family of 800 ms square voltage pulses from a holding potential of
70 mV to a range of potentials between
80 and þ40 mV (10 mV increments). The lower panel shows the activation curve from the current traces shown above. Plotted are the tail current amplitudes versus the test-pulse voltage. The tail-current amplitudes reflect the fractional activation of the channels at the end of the preceding test pulses. (b) Superimposed are the current-clamp record from an SCG neuron (black) and the voltage-clamp trace of Kv7.2/7.3 current activation from a CHO cell expressing the channels (red). Both traces are synchronized for timescale. (See text for full caption.)

FIGURE 11.2 Gq/11-coupled signaling in mammalian cells. Depicted is the canonical pathway triggered by the activation of Gq- or G11-coupled receptors in neurons: activation of phospholipase C (PLC) results in hydrolysis of membrane phosphatidylinositol 4,5- bisphosphate (PIP2, brown shapes) and release of inositol triphosphate (IP3, brown octagons) and diacylglycerol (DAG, green rectangle). The former may trigger release of Ca2þ (green spheres) from IP3-sensitive stores, whereas the latter can activate protein kinase C (PKC, ochre shape) or be further degraded to phosphatidic (PA, pink ovals)or arachidonic (AA, green ovals)acids.Each of these intermediates is capable of triggering diverse downstream signaling pathways.

FIGURE 11.4 Purinergic stimulation provokes PIP2 hydrolysis, but does not depress ICa. (a) Plotted are normalized Ca2þ current amplitudes recorded from a superior cervical ganglion (SCG) neuron under perforated patch-voltage-clamp. (See text for full caption.)

Slo1 (KCNMA1) gene (a)

* **

5'-UTR

1

2

*

3 4 5

6 7

8

* *

*

9 10 11 12 13 14 15 16 17 18 19 20 21

22 23 24 25

* 26

27 3'-UTR TGA

ATG

400 bp

M1 M2 M3 M4 mSlo1 MANGGGGGGGSSGGGGGGGGGSGLRMSSNIHANHLSLDASSSSSSSSSSSSSSSSSSSSSVHEPKMDALIIPVTMEVP hSlo1 MANGGGGGGGSSGGGGGGGG-SSLRMSSNIHANHLSLDASSSSSSSSSSSSSSSSSSSSSSVHEPKMDALIIPVTMEVP

(b)

*

N

3,4

+

+ S5 S1 S2 S3 S4

S0

+

2,3

* * 1,2

SSQA(V)DG RCK1

S6 DRD(G)DV

9,10

*

18,19

*

*

16,17

S7

S8

S10

S9

C RCK2 23,24

*

* “Ca2+ bowl”

FIGURE 12.1 Gene map and protein topology of Slo1 (KCNMA1) gene. (a) Gene map showing constitutive exons (gray boxes, at scale) and introns (broken lines, not to scale) of human Slo1. Known splice sites in various species are marked with an arrow and a star. Enlarged region shows probable translation start sites (methionine, M1–M4) in human and mouse and high degree of homology between species, which applies for the rest of the protein (not shown). Most of studies have used clones beginning at M3 (red). (b) Slo1 protein topology. Seven transmembrane domains are marked as S0–S6. Cytoplasmic hydrophobic domains are marked as S7–S10. Regulator of conductance of Kþ domains, RCK1 (blue) and RCK2 (pink) positions (T. Yusifov and R. Olcese, personal communication). Ca2þ bowl, calcium sensing domain (green). Arrowheads mark junctions of translated constitutive exons; numbers of corresponding exons are only given for known alternative splice sites (*) (e.g., 1,2 marks where the translated protein of exon 1 joins the translated protein of exon 2). Red boxes and sequences mark human mutations linked to generalized epilepsy and paroxysmal movement disorder (D/G), and autism (A/V).

FIGURE 12.3 A simplified scheme of Slo1 channel transcriptional regulation by estrogen, channel maturation, and degradation. Estrogen binds to the estrogen receptor a after crossing the plasma membrane, causing dimerization of the receptor. Dimerized receptor enters the nucleus and binds to estrogen response elements (ERE) present in Slo1 gene enhancing transcription. mRNA migrates to the cytoplasm and interacts with the translational machinery to form polyribosomes. Slo1 protein is inserted to endoplasmic reticulum (ER) membrane where it assembles into tetramers and associates or not associates with b subunits (depending on the cell type). Tetrameric channels are transferred to the cell membrane. Splice variant 1 insert (SV1) retains insertless Slo1 and b1 subunit in the ER. Surface expression of the channel can be regulated by b subunits via endocytosis. Direct binding of estrogen to Slo1 induces channel degradation by the proteasome. Dotted arrows indicate probable events: phosphorylation favoring forward traffic and association with partners in the cytosol during traffic to the membrane.

FIGURE 13.2 The overall architecture of the KChIP1–Kv4.3N complex. All three panels have the same color codes with some secondary structural elements labeled specifically. (a) One KChIP1 molecule in gold interacts simultaneously with two Kv4.3Ns in blue. (b) The 4:4 complex of KChIP1–Kv4.3N shown on this panel is generated from the complex on panel A through symmetric operations. (c) KChIP1–Kv4.3N complex in 4:4 showing the clamping effect of KChIP1 molecule on the tetramer of Kv4.3. (See text for full caption.)

FIGURE 13.3 Significant structural rearrangements occur to KChIP1 upon Kv4.3 N-terminal binding. Surface representations are shown to an isolated KChIP1. (See text for full caption.)

FIGURE 13.4 The N-terminal inactivation peptide of Kv4.3 is completely sequestered in a hydrophobic groove on the surface of KChIP1. (a) The N-terminal hydrophobic peptide of Kv4.3 binds to an elongated hydrophobic groove on the surface of KChIP1. KChIP1 is shown in the surface representation, and the N-terminal inactivation peptide of Kv4.3 is shown in ribbon. The Kv4.3 N-terminal residues involved in hydrophobic contacts with KChIP1 are colored in magenta. (b) An SDS-PAGE gel showing the pull-down results for the point mutants of both KChIP1 and the N-terminal peptide of Kv4.3. The single mutation of KChIP1 Y134E and the triple mutation W8E, P10E, and A15E in Kv4.3 completely abolish their interactions with their respective wild-type partners. GST-pull-down assay is used to detect the interaction between WT Kv4.3N and KChIP1 mutants (left panel), whereas various His-tagged Kv.3 mutants are immobilized on Ni-resin and WT KChIP1 is allowed to flow through Ni-resin (right panel). In both cases, the resin is visualized on SDS-PAGE followed by coomassie staining. (c) Sequence alignments of the N-terminal peptide of Kv4-family proteins among different species. (d) Effects of KChIP mutation within the first interface on Kv4.3. The representative traces recorded from oocytes injected with cRNAs of WT Kv4.3 alone, WT Kv4.3 þ WT KChIP1, and WT Kv4.3 þ KChIP Y134E. The left side panel depicts currents recorded from oocytes held at
80 mV by a family of pulses from
60 to 40 mV in 10 mV increments for 1 s, and the right panel shows recovery from inactivation for varying lengths of time at step from
80 to þ40 mV. (e) The representative traces of Kv4.3 triple mutant (W8E-P10E-A15E) alone and the triple mutant þ WT KChIP1 recorded from oocytes under the same protocol as in (d).

FIGURE 13.5 The second contact interface between KChIP1 and Kv4.3 T1 domain. (a) The interaction within the second interface is mediated by both hydrophobic contacts and salt bridges. The KChIP1 molecule and the Kv4.3 peptide (residues 70–78) are shown in surface representation and dark green ribbon, respectively. The residues of Kv4.3 colored in magenta are involved in hydrophobic interaction with KChIP1. The blue, red, and white represent the positive, negative, and hydrophobic surface of Kv4.3, respectively. (b) A close-up view of the interaction within the second interface between KChIP1 and Kv4.3N. The KChIP1 molecule is shown in light green ribbon and colored in green, and the residues involved in the interaction with Kv4.3N are highlighted in yellow. The O and N atoms are shown as red and blue, respectively. The salt bridges are indicated by black dashed dots. (c) Sequence alignment of Kv4.3 peptide (70–78) with corresponding sequences of other Kv proteins. The residues F73 and F74 of Kv4-family labeled in magenta square on the top form hydrophobic contacts with KChIP1 molecule and those with blue squares make salt bridges with KChIP1. (d) The representative traces are recorded from oocytes injected with cRNAs of Kv4.3 double mutant (E70A, F73E) that disrupts the second interface, in the presence or absence of WT KChIP1 or the KChIP1 the triple mutant (L39E, Y57A, K61A). The left panel depicts currents from oocytes held at
80 mV by a family of pulses from
60 to 40 mV in 10 mV increments for 1 s, and the right panel shows recovery from inactivation for varying lengths of time at step from
80 to þ40 mV every 8 s.

FIGURE 13.7 Comparison of the modeled Kv4.3–KChIP1 channel complex with Kv1.2– Kvb2. (a) Side views of Kv1.2/Kv4.3 T1-KChIP complex in which Kv4.3 T1 domain fused with transmembrane-spanning domains of Kv1.2 (left panel) and Kv1.2-Kvb2 complex (right panel). The tetrameric subunits of Kv1.2 channels are labeled with colors of cyan, yellow, pink, and green, respectively. KChIP1 and Kvb2 are labeled in blue and wheat, respectively. (b) Top views of panel A, showing KChIPs positioned between two adjacent T1 domains (left panel) and Kvb2 beneath the T1 domains (right panel).

PART I NEURONAL VOLTAGE-GATED ION CHANNEL FUNCTIONS

1 NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS ALEXANDER SCRIABINE1 AND DAVID J. TRIGGLE2 1

Department of Pharmacology, Yale University School of Medicine, New Haven, CT 06520, USA 2 School of Pharmacy and Pharmaceutical Sciences, State University of New York, Buffalo, NY 14260, USA

1.1 STRUCTURE AND DISTRIBUTION Although the L-type calcium (Ca2þ) channel has long been associated with cardiovascular physiology and pharmacology and the pharmacological and therapeutic effects of a structurally diverse group of blockers, notably of the 1,4-dihydropyridine family (Fleckenstein, 1983; Goldmann and Stoltefuss, 1991; Triggle, 2004, 2006) (Fig. 1.1), it is also widely distributed in the peripheral and central nervous system where its roles are being increasingly examined. The application of established calcium channel antagonists, such as nifedipine, does not appear to have dramatic neuronal effects, but application of dihydropyridine activators, such as Bay K 8644, produces profound neuronal and behavioral disturbances, indicating the potential pathological and therapeutic importance of these channels (Lipscombe et al., 2004; Striessnig et al., 2006). The apparent general lack of the effect of 1,4-dihydropyridines on transmitter release suggests that L-type channels play an unimportant role in presynaptic calcium entry coupled to transmitter release, but rather are involved in longer term events such as neuronal plasticity and the control of gene expression.

Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

3

4

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

FIGURE 1.1 The structural formula and receptor site organization for the three principal drug classes active at the L-type calcium channel. A separate receptor site exists for each of the drug classes. That for 1,4-dihydropyridine activator and antagonists has been best described and explored.

However, this may be an oversimplification based in part upon an excessive reliance on the prototypical pharmacology of 1,4-dihydropyridines to define channel properties and function. The neuronal L-type voltage-gated calcium channel (NLTCC) is a member of a family of voltage-gated calcium channels, which in turn belong to a superfamily of voltage-gated ion channels, including those for potassium (Kþ) and sodium (Naþ). It is likely that the potassium channel represents an ancestral member whose functions have been progressively modified by processes of gene duplication and mutation. A number of general reviews are available for this superfamily of ion channels (McDonough, 2004; Catterall, 2005; Lacinova, 2005; Zamponi, 2005; Triggle, 2006). The voltagegated calcium channel family is a heteromeric association of subunits as depicted in Fig. 1.2—a1, b, a2d, and g—and their biophysical and pharmacological properties as well as their expression are influenced significantly by the totality of the subunit interactions. Since there are several members of each subunit class with alternative splicing, the number of potential permutations with attendant variations in localization and biophysical and pharmacological properties is quite large. 1.1.1

Classification

Voltage-gated calcium channels are divided into two main classes, high voltage activated (HVA) and low voltage activated (LVA), and each of these classes is further subdivided (Catterall et al., 2005). The HVA channels are subdivided into five types (L, T, N, P/Q, and R) on the basis of their physiological and pharmacological properties. An interesting account of the history of the discovery of calcium channels has been provided by Tsien and Barrett (2005). The properties of L-type (for longlasting) channels are summarized in Table 1.1. Figure 1.3 depicts the overall sequence

STRUCTURE AND DISTRIBUTION

TABLE 1.1

5

Properties of L-Type Voltage-Gated Calcium Channels (Cav1.1–1.4)

Physiological properties Conductance (pS) Activation threshold Deactivation rate Inactivation rate Permeation Function

25 High Fast Slow Ba2þ > Ca2þ E–C coupling, CV system, smooth muscle, endocrine cells, neurotransmitter release (eye, ear)

Pharmacological properties 1,4-Dihydropyridines Phenylalkylamines Benzothiazepines Benzimidazoles v-Conotoxin GV1A v-Conotoxin MVIIC v-Agatoxin IVA v-Agatoxin IIIA Cd2þ block Ni2þ block Radioligands of choice Tissue expression Disease relevance

Sensitive Sensitive Sensitive Insensitive Insensitive Insensitive Insensitive Sensitive Potent Weak 3 ½H
-cis-(þ)-Diltiazem, 3 ½H
-desmethoxyverapamil, 3 ½H
-isradipine Widespread: CV system, neurons, endocrine tissue, skeletal muscle, smooth muscle Hypertension, angina, malignant hyperthermia, hypokalemic periodic paralysis, night blindness

FIGURE 1.2 A schematic representation of the organization of the subunits of the voltagegated calcium channel.

6

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

FIGURE 1.3 The organization of the voltage-gated calcium channel family.

homology between the several channel classes. The L-type voltage-gated channel, the principal focus of this chapter, belongs to the HVA class (although the potential dependence of activation and inactivation varies between subtypes) and has been particularly well investigated from structural, functional, and pharmacological perspectives. Drugs acting at this class of channel (Fig. 1.1) not only have found widespread use in the treatment of a number of cardiovascular disorders, but have also been widely explored for their potential application in a number of other disorders, including neuronal pathologies, from achalasia through depression to tinnitus and vertigo. 1.1.2

Structure

A schematic representation of the overall organization of the voltage-gated calcium channel a1 subunit is depicted in Fig. 1.1. The a1 subunit consists of four homologous domains, and the S4 segments contain regularly arrayed positively charged lysine and arginine residues, a feature associated with the voltage sensitivity and channel opening properties of these channels. This subunit makes up the pore-forming and voltagesensing components of the channel as well as containing the major drug binding sites. The S5–S6 linkers each contain a critical glutamic acid residue that comprises in total the selectivity filter of the channel. A summary of the sizes of individual cloned a1 subunits of the Cav1.1–1.4 (L-type) family is provided in Table 1.2. Interaction with other subunits is important for both expression and biophysical and pharmacological properties of the channel.

STRUCTURE AND DISTRIBUTION

TABLE 1.2

7

Sizes of Cloned Cav1 a1 Subunits

Subunit

Origin

MW (kDa)

No. of Residues

Cav1.1 Cav1.2 Cav1.2 Cav1.2 Cav1.3 Cav1.4

Rabbit skeletal muscle Rabbit heart Rabbit lung Rat aorta Human pancreas Human retina

212 242.8 242.5 243.6 247.6 219.5

1873 2171 2166 2169 2181 1966

Data from compilation of Lacinova (2005).

The cytosolic b subunit b1–4, coded by four genes (CACNB1–4) and with a number of splice variants, interacts through specific domains on that subunit and on the S1–S2 linker: the b interaction domain (BID) of approximately 30 residues and the a interaction domain of approximately 18 residues (Pragnell et al., 1994; Hofmann et al., 1999; Van Petergem et al., 2004; Cens et al., 2005; Doering and Zamponi, 2006). The b subunits are widely distributed in excitable tissues. The b subunit resembles a membrane-associated guanylate kinase, the GK domain of which provides a hydrophobic cleft for calcium channel binding (Takahashi et al., 2004). The AID–BID complex may also provide an additional site for drug interaction (Triggle, 2004, 2006). The a2d 1–4 subunit, coded by four genes CACNA2D1–D4 and with a number of splice variants, comprises two components linked by a disulfide bond with a membrane-spanning d component and an extracellular a2 component (De Jongh et al., 1990; Doering and Zamponi, 2006). These subunits are also widely distributed in excitable tissues. The a2d4 subunit also generates a drug binding site, characterized in particular for drugs such as gabapentin and pregabalin used for pain relief. The g 1–8 subunit is coded by eight genes, CACNG1–8, and is of particular importance in skeletal muscle (Cav1.1) and neuronal (Cav2 and Cav3) channels (Lacinova, 2005; Doering and Zamponi, 2006). The g subunits are integral membrane proteins with four transmembrane domains and intracellular C and N termini. Of particular importance to understanding both channel function and pharmacology is the question of the actual subunit combinations that make up the various channel types: it is likely that this is tissue and cell specific and the details remain largely unresolved. 1.1.3

Subtypes

Four principal classes of the L-type channel exist represented by the a subunit of Cav1.1–1.4 and associated with the b, a2d, and particularly for the Cav1.1 class, the g subunit. The specific permutations of association of these subunits and their splice variants remain to be fully established. However, the Cav1.1 channel, which mediates excitation–contraction coupling in skeletal muscle, is composed of the principal a1 subunit together with b1, a2d1, and g 1 subunits. The organization of the other three L channel types remains less certain, and it is possible that the subunit compositionvaries and is tissue dependent (Cens et al., 2005; Doering and Zamponi, 2006). In addition to the primary classification, other subtypes of the channel may exist through the

8

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

TABLE 1.3 Channel

Interaction of 1,4-Dihydropyridines with Splice Variants of the L-Type

Nifedipine Nisoldipine (þ)-Isradipine SDZ 2017-180

KD (nm), 100 mV, CaV1.2a (Cardiac)

KD (nm), 50 mV, CaV1.2b (Smooth Muscle)

47 2.1 15 91

10 0.56 2.1 100

Data from Morel et al. (1998).

existence of splice variants. Such subtypes may exhibit both tissue-dependent localization and tissue-selective pharmacology (Soldatov et al., 1986; Welling et al., 1993, 1997; Hu and Marban, 1998; Morel et al., 1998; Zhulke et al., 1998; Lacinova et al., 2000; Safat et al., 2001). Thus, several comparisons have been made of the properties and 1,4-dihydropyridine sensitivity of splice variants of the Cav1.2 (cardiac and smooth muscle) subunit. The a-splice variant (cardiac isoform) has a lower sensitivity to 1,4-dihydropyridines than the b-splice variant (smooth muscle). Representative data are shown in Table 1.3. Splice variants have been investigated in other tissues, including brain and neuroendocrine cells, but evidence for selective distribution and function is lacking (Safat et al., 2001). 1.1.4

Distribution

Although the dominant physiological and pharmacological expression of L-type channel activity is usually assumed to be localized to the cardiovascular system, these channels are in fact widely distributed throughout the body including the peripheral and central nervous systems (Bean, 1989; Hess, 1990; Miljanich and Ramachandran, 1995; Bech-Hansen et al., 1998; Strom et al., 1998; McRory et al., 2004; Herlitze and Mark, 2005; Lai and Jan, 2006). Within neurons, selective cellular localization occurs. Cav1.4 is present in the retina where loss-of-function mutations cause night blindness (Bech-Hansen et al., 1998; Strom et al., 1998; McRory et al., 2004). Cav1.2 and Cav1.3 are more widely distributed in nervous tissue, neuroendocrine cells, and hair cells (Herlitze and Mark, 2005). The role of Cav1.3 channels in hair cells in the cochlea is linked to the development of these cells and to the associated development of highconductance calcium-activated potassium BK channels (Nemzou et al., 2006). The widespread distribution of the Cav1.2 and Cav1a.3 channels within both the cardiovascular and nervous systems has made determination of their neuronal roles through pharmacological intervention difficult since the available L-type channel ligands have powerful cardiovascular properties that may overshadow or complicate any activities produced in neurons. However, selective elimination of 1,4-dihydropyridine sensitivity from Cav1.2a1 subunits permits the role of Cav1.3 channels to be examined through pharmacological dissection (Bourient et al., 2004). Cav1.3 stimulation was shown to selectively contribute to Fos expression, to neurotransmitter release in the ventral striatum, and to be associated with depression-like behavioral effects.

STRUCTURE AND DISTRIBUTION

9

Within neurons, L-type channels enjoy selective localization. In rat cortex and hippocampus, Cav1.2 and Cav1.3 channels are principally localized in cell bodies and proximal dendrites, with the Cav1.2 type being concentrated in clusters and the Cav1.3 more dispersed (Hell et al., 1993; Ludwig et al., 1997). In apparent contrast, L-type channels are associated in rat globus pallidus neurons predominantly with distal dendrites, where the authors suggest that the proximal localization reported in earlier studies may represent channels in the process of transport (Hanson and Smith, 2002). The issue of the subunit association of these a1 subunits remains unclear, but the work of Ludwig et al. (1997) on the rat brain suggests that this may be cell specific, thus providing further diversity of channel structure and function. NLTCCs do not have a major role in neurotransmitter release but are certainly involved in the control of gene transcription activity (Dolmetsch et al., 2001; Zhang et al., 2002; Deisseroth et al., 2003; Evans and Zamponi, 2006), and the subsequent changes in protein expression may be linked to changes in synaptic strength and the regulation of transmitter phenotype (Brosenitsch et al., 1998; Deisseroth et al., 2003). Proteolytic cleavage of the Cav1.2 channel generates a C-terminal fragment, calcium channel-associated transcription regulator (CCAT), that translocates to the nucleus where it interacts with an endogenous promoter to control the expression of a number of genes associated with signaling and excitability in neurons (Gomez-Ospina et al., 2006). Other voltage-gated calcium channels behave similarly, and this may be a general control mechanism (inter alia, Hell et al., 1993; Westenbroek et al., 1998; Kordasiewicz et al., 2006). However, the level of such activity may differ between L-type channel subtypes: in rat hippocampal neurons, Cav1.3 plays a more important role in pCREB signaling than does Cav1.2 (Zhang et al., 2005b, 2006). 1.1.5

Mechanisms of Action: Activation, Inactivation, and Drug Action

The Cav1.1–1.4 class of channels show overall similar structure and pharmacology but differ quantitatively in a number of important aspects. The traditional view of this channel class is that they are activated by strong depolarization, are relatively slowly activated, have large single-channel conductance, show calcium-dependent inactivation, and are very sensitive to the 1,4-dihydropyridine family of ligands (Lipscombe et al., 2004). However, at least some channels formally of the Cav1.2 and 1.3 classes, those most widely distributed in the nervous system, show distinct behavior, in particular activating at relatively polarized levels of membrane potential and showing differential sensitivity to 1,4-dihydropyridines (inter alia, Xu and Lipscombe, 2001; Lipscombe, 2002; Lipscombe et al., 2004; Helton et al., 2005). Recombinant neuronal Cav1.2 and Cav1.3 channels open rapidly over a wide range of membrane potentials and carry significant calcium current in response to single stimulus. However, 1,4-dihydropyridines do not block calcium entry in response to single action potential stimuli but are effective in blocking current in response to step depolarization or to long trains of action potentials. This reflects the significant state-dependent interactions of 1,4-dihydropyridines (Section 1.2.2). Furthermore, 1,4-dihydropyridines completely block Cav1.2 channels, but only partially block Cav1.3 channels (Helton et al., 2005). These observations indicate that the role of neuronal L-type channel activation in response to brief stimuli may have been underestimated because of an excessive

10

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

reliance on 1,4-dihydropyridines as pharmacological markers of L-type channel activity. 1.1.6

Interaction with Other Cellular Components

In addition to interaction with b, g, and a2d subunits, the principal a1 subunit of the voltage-gated calcium channel interacts with a variety of other proteins that regulate its expression, trafficking, and activity (for reviews see Catterall et al., 2005; Herlitze and Mark, 2005; Lee and Catterall, 2005; Stanley and Chan, 2005; Evans and Zamponi, 2006). Channels of the Cav2.1 and 2.2 classes interact with the bg subunits of G proteins to mediate differential inhibition and with the SNARE protein complex that mediates exocytosis of transmitters. Cav1.1 channels interact with ryanodine receptors to mediate excitation contraction coupling in skeletal muscle and also with A kinase anchoring proteins (AKAPs) that anchor cAMP-dependent protein kinase. Cav1-type channels undergo both Ca2þ-dependent inactivation and facilitation, best established for Cav1.2 channels, mediated predominantly through calmodulin binding to the IQ-domain of the C-terminal portion of the a1 subunit (reviewed in Lee and Catterall, 2005).

1.2 CLASSES OF DRUGS Of the three structural classes of drugs depicted in Fig. 1.1—the benzothiazepinones, the phenylalkylamines, and the 1,4-dihydropyridines—the latter represent the largest class studied both clinically and experimentally. They are also quantitatively the most active, exhibiting pharmacology in the nanomolar concentration range, and they also include both antagonist and activator species. However, drugs active at L-type channels are not confined to these three structural scaffolds, and many diverse agents exhibit antagonist properties. The structure–function relationships of drugs active at L-type channels have been extensively reviewed over many years (inter alia, Janis and Triggle, 1983, 1984a, 1984b; Janis et al., 1987; Triggle et al., 1989; Goldmann and Stoltefuss, 1991; Rampe and Triggle, 1993; Triggle, 2003, 2004, 2006; Budriesi et al., 2007). Hence, only a very brief overview primarily of the 1,4-dihydropyridines, the principal therapeutic and molecular tools active at the L-type channel, will be presented here. 1.2.1

Structural Requirements

The basic structural requirements for antagonism and activation are depicted in Fig. 1.4, and the structural formulas of clinically available 1,4-dihydropyridines are depicted in Fig. 1.5. The 1,4-dihydropyridines are of particular interest for several reasons. First, they are extremely potent ligands for the L-type channel. Second, they exhibit both activator and antagonist properties. Third, they exhibit considerable stereoselectivity of action. Fourth, they show substantial state-dependent mode of interaction. Fifth, the 4-aryl-1,4-dihydropyridine nucleus is a “privileged structure’’ capable, when decorated with the appropriate substituents, of interacting with a variety of receptors and ion channels (Triggle, 2003).

CLASSES OF DRUGS

11

FIGURE 1.4 The structural features that determine activity of activator and antagonist in the 1,4-dihydropyridine family of drugs active at the L-type calcium channel.

Bay K 8644 serves as the prototypical activator and such compounds show highly differential pharmacology whereby one enantiomer is an activator and the other an antagonist (Fig. 1.6). Stereoselectivity is quite generally observed with all 1,4-dihydropyridines, but the extent of chirality depends on the substituents around the 1,4-dihydropyridine ring. Thus, nitrendipine has a modest stereoselectivity factor of approximately 5–10, whereas amlodipine has a (+) factor of approximately 1000.

FIGURE 1.5

The clinically available 1,4-dihydropyridines.

12

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

FIGURE 1.6 Stereochemical requirements for activator and antagonist activity in 1,4dihydropyridines. The S-enantiomers are activators and the R-enantiomers are antagonists.

1.2.2

State-Dependent Actions

The 1,4-dihydropyridines (as well as verapamil and diltiazem) are characterized by their state-dependent mode of interaction at the L-type channel. These interactions arise because drugs may exhibit differential affinity and/or access to their binding sites in the resting, open, or inactivated states of the channel (Hille, 1977; Hondeghem and Katzung, 1985; Triggle, 1989; McDonough and Bean, 2006). Transitions between these states are determined by changes in chemical or electrical potential and by the kinetics of channel opening and closing. Drugs that interact preferentially with the open or inactivated state of the channel will show an increase in apparent affinity under those physiological or pathological conditions that increase the availability of those channel states. In addition, the structure of the drug and overall physicochemical properties—charged/uncharged and hydrophilic/hydrophobic characteristics—can control the access of the drug to the receptor site. Charged and polar drugs may access their binding sites through a polar and hydrophilic pathway, including the channel pore, whereas nonpolar drugs can access binding sites through membranedelineated pathways. Both diltiazem and verapamil exhibit frequency-dependent interactions and hence their use, particularly of verapamil, in certain tachyarrhythmias, whereas nifedipine and other 1,4-dihydropyridines exhibit voltage-dependent interactions consistent with a preferential interaction with the inactivated states of the channel, a property that underscores their general vascular selectivity (Bean, 1984; Sanguinetti and Kass, 1984; Wei et al., 1986; Triggle, 1989; Zhen et al., 1992; Sun and Triggle, 1995).

FUNCTION

1.2.3

13

Privileged Structures

Finally, the 4-aryl-1,4-dihydropyridine nucleus is a privileged structure and with appropriate ring substituents can access a diverse set of channels and receptors (Triggle, 2003). Certain dihydropyridines also interact with T-, N-, and P/Q-type calcium channels (Cohen et al., 1992; Kumar et al., 2002; Zhou et al., 2002; Yamamoto et al., 2006). Agents such as amlodipine, cilnidipine, barnidipine, benidipine, and nicardipine that can also block N-type channels may have some therapeutic cardiovascular advantage as well as interact at the N-type channels in neuronal tissues.

1.3 FUNCTION 1.3.1

Role of Subunits and Isoforms

The role of NLTCCs in the neuronal function is determined by the type and location of neurons as well as by the composition of channels, their subunits, and isoforms. Each channel complex consists of the pore-forming a1 subunit and three regulatory subunits (a2, b, and g). Three isoforms of the a1 subunit have been identified in the central nervous system: Cav1.2a1, Cav1.3a1, and Cav1.4a1. They have distinctly different neurological functions. The first two isoforms often occur together, expressed in the same cells, while Cav1.4a1 is found mainly in the retinal neurons (Baumann et al., 2004). The regulatory subunits also contribute to heterogeneity of NLTCCs. The a2d1 subunit has been intensively studied and identified as a molecular target for the analgesic action of pregabalin and gabapentin (Field et al., 2006; Joshi and Taylor, 2006). The techniques available to study physiological functions of NLTCCs are either pharmacologic or genetic. Pharmacologic methodology calls for the use of L-type Ca2þ channel activators or antagonists. Unfortunately, specific ligands for isoforms of the a1 subunit of NLTCCs have not yet been discovered, and the currently available ligands for L-type channels are not isoform specific. The genetic methodology utilizes “knockout’’ mice lacking specific isoforms of NLTCCs (Schulla et al., 2003; Sinnegger-Brauns et al., 2004). The neurological phenotype of Cav1.3a1/ mice exhibits inner hair cell dysfunction and cochlear sensory cell degeneration but appears neurologically normal. It has been suggested (Clark et al., 2003) that certain Ca2þ channel isoforms may support distinct behavioral functions. New insights into the role of NLTCCs and the Cav1.3a1 isoform in neuronal function were provided by the discovery of a link between these channels and macromolecular signaling complex formed by Shank and other modular adapter proteins as well as a link with G-proteincoupled receptors (Olson et al., 2005; Zhang et al., 2005). Using a mouse model without dihydropyridine (DHP)-sensitive Cav1.2a1 subunits (Cav1.2 DHP/ mice), Sinnegger-Brauns et al. (2004) found that in the ventral striatum of these mice agonistinduced glutamate and 5-HT release was abolished, while dopamine and norepinephrine release remained intact. This observation demonstrated differences in the functions

14

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

of Cav1.2a1 and Cav1.3a1 isoforms, although both isoforms appear to control emotional behavior in mice. On the basis of experiments suitable for the dissection of function of the two isoforms, Striessnig et al. (2006) concluded that selective inhibitors of channels containing Cav1.3a1 isoforms can be expected to have antidepressant and anxiolytic properties. 1.3.2

Synaptic Plasticity and Memory

A brief period of high-frequency electrical activity applied artificially to a neuronal pathway can enhance the strength of synapses for various periods of time. This phenomenon is called long-term potentiation (LTP). It can be induced in the cerebral cortex, hippocampus, and other brain areas. Many features of LTP resemble those involved in memory storage, and LTP is widely used in memory research in attempts to elucidate molecular mechanisms of memory (Kandel, 2001). In rat hippocampus, there are at least three types of LTP (short lasting or LTP1, of intermediate duration or LTP2, and long lasting or LTP3). LTP3 is selectively dependent on NLTCCs (Raymond and Redman, 2006). In rat basolateral amygdala, LTP is induced at least in part due to influx of Ca2þ through channels containing Cav1.2 isoforms (Pinard et al., 2005). In the superior cervical ganglion of the rat, however, ganglionic transmission is mediated primarily by P/Q- and N-type channels with only 14% contribution by L-type calcium channels (Cifuentes et al., 2004). Antagonists of NLTCCs, for example, nimodipine, reduce the extent of LTP in rat hippocampal neurons and abolish the induction of long-term depression (LTD), induced by postsynaptic spiking prior to presynaptic activation (Bi and Poo, 1998). The role of hippocampal NLTCCs containing Cav1.2 channels in synaptic plasticity and spatial memory was studied in Cav1.2HCKO mice in which the CACNAIC (Cav1.2) gene was inactivated (Moosmang et al., 2005b). These investigators found that the late phase of long-term potentiation (L-LTP) is lost in the hippocampus and neocortex of these animals and hippocampus-dependent spatial memory is severely impaired. A decreased activation of mitogen-activated protein kinase (MAPK) pathway and a reduced cAMP response element-dependent transcription were found in CA1 pyramidal neurons of Cav1.2HCKO mice. Phosphorylation of cAMP response element binding protein (CREB) at Ser133 is considered to be an important step in the induction of gene expression critical for memory (Moosmang et al., 2005a); it is impaired in Cav1.2HCKO mice. These observations suggest that selective inhibitors of Cav1.2 subtypes of NLTCCs could impair spatial memory. On the contrary, there is evidence that calcineurin, the only Ca2þ-activated protein phosphatase in the brain, negatively modulates learning, memory, and neuronal plasticity (Mansuy, 2003). Calcineurin has been identified as the key signal in the extinction of fear memory (Lin et al., 2003). It also impairs spatial memory in mice (Mansuy et al., 1998). Ca2þ needed for activation of calcineurin enters neurons at least partially through NLTCCs and is likely to enhance calcineurin-induced negative modulation of learning and memory, so that inhibitors of NLTCCs could be expected to improve learning and spatial memory. It is currently unknown whether Ca2þ entry through channels containing Cav1.2 or Cav1.3 isoforms is linked to the activation

FUNCTION

15

of calcineurin. If it is, a selective inhibitor of these isoforms could improve learning and spatial memory. The idea that antagonists of NLTCCs could improve memory and learning and be useful in the treatment of Alzheimer’s disease was originally based on the calcium hypothesis of Alzheimer’s disease and aging (Landfield, 1987; Disterhoft et al., 1994; Khachaturian, 1995). One of the key elements of this hypothesis involved breakdown of [Ca2þ] homeostasis and elevation of intraneuronal calcium as a factor contributing to neuronal degeneration and death. An antagonist of NLTCCs, nimodipine, has been shown to facilitate learned behavior in rats with neocortical injury (LeVere and Sandin, 1989). More recently, nimodipine has been shown to ameliorate age-related memory decline in aged rats. This effect was associated with the decline in abnormally high level of expression of channels containing Cav1.3 subunit in the hippocampus of these animals (Veng et al., 2003). Some clinical studies suggested that nimodipine activates cognition in patients with vascular or primary degenerative dementia (Tobares et al., 1989; Fischhof et al., 1993) and another antagonist nitrendipine (Forette et al., 1998) reduces the incidence of dementia in the elderly hypertensive population. High cytosolic calcium concentrations in neurons inhibit asecretase cleavage of amyloid precursor protein (APP) and increase intraneuronal levels of b-amyloid peptide (Ab1–42) (Pierrot et al., 2004). NLTCCs antagonists can be expected to reduce cytosolic calcium in neurons. Morich et al. (1996) reported clinical experience with nimodipine in patients with probable Alzheimer’s disease (AD). There was no convincing evidence of effectiveness of nimodipine in AD, but this drug improved performance of patients in Buschke’s Selective Reminding test, which is considered to be an index of memory storage. This finding suggests that Ca2þ entry through NLTCCs may modulate storage of memory in humans. The failure to demonstrate convincingly cognition activation with antagonists of NLTCCs in humans may be explained by the dual role of intraneuronal Ca2þ. It improves some aspects of memory but facilitates neurodegeneration and activates calcineurin, which impairs cognition. Antagonists of NLTCCs may also have effects other than those mediated by calcium channels, and these effects may oppose the consequences of calcium channel blockade. Nimodipine, for example, has been reported to stimulate Ab1–42 secretion in neuroblastoma cell cultures, an effect apparently not involving calcium channels (Facchinetti et al., 2006). This effect, if it occurs in vivo, would tend to oppose the putative cognition enhancing effect of this drug. 1.3.3

Pain

During the last two decades, substantial evidence has accumulated that voltage-gated calcium channels (VGCCs) are involved in the perception of pain (Cao, 2006; Yaksh, 2006). Pain behavior appears to be altered primarily by the Cav2.2 subunits of N-type channels, located in the presynaptic terminals where they seem to control neurotransmitter release (e.g., glutamate or substance P). The involvement of calcium channels in the control of pain was supported by the discovery of the analgesic activity of a2d subunit ligands, gabapentin, pregabalin, and L-phenylglycine, in neuropathic pain (Taylor, 2004; Frampton and Foster, 2005; Lynch et al., 2006), although not all ligands of this subunit attenuated neuropathic pain in rat spinal cord ligation model

16

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

(Lynch et al., 2006). The a2d protein is encoded by four genes, a2d1, a2d2, a3d 3, and a2d4, and replacement of a single amino acid (arginine in a position 217) in a2d1 subunit prevented gabapentin or pregabalin binding and nearly abolished their analgesic activity in mutant mice (Field et al., 2006). This finding strongly supports the importance of the a2d 1 subunit for the analgesic action of these two compounds. The involvement of NLTCCs in pain perception is likely to depend on the nature of the a2d1 subunit in these channels. Some of the early pharmacologic studies with L-type calcium channel antagonists, for example, 1,4-dihydropyridines, demonstrated their antinociceptive activity and interactions with opiates (Hoffmeister and Tettenborn, 1986; Del Pozo et al., 1987). The activity was dependent on the test, doses of the drugs, and routes of their administration, and some of the findings were contradictory. Nimodipine and nifedipine antagonized acetic acid-induced writhing following their intracerebroventricular (i.c.v.) administration to mice (Miranda et al., 1993). In the tail withdrawal test, the same drugs had antinociceptive activity in rats by chronic subcutaneous (s.c.) administration (Martin et al., 1996). In hot plate test, however, nimodipine, but not nifedipine, verapamil, or diltiazem, had analgesic effect (Miranda et al., 1992). It has been suggested that 1,4-dihydropyridines may affect pain perception at the spinal level (Martin et al., 1996). On the basis of the available evidence, L-type calcium currents appear to be only marginally involved in pain perception. The possibility that antagonists of NLTCCs could be useful as adjuncts to analgesics has been extensively studied. Many experimental studies explored the interaction of 1,4-dihydropyridines with opiates. In the rat tail-flick test, nimodipine, at 1 mg/kg i.p., or lercanidipine, at 0.3 mg/kg i.p., potentiated analgesia caused by k-opioid receptor agonists and prevented the development of tolerance to the opiates (Gullapalli and Ramarao, 2002a). Chronic administration of nimodipine to rats at 1 mg/kg/day i.p. for 10 days increased morphine (2 mg/kg i.p.) induced analgesia. The effect was additive to that of naloxone (Gullapalli and Ramarao, 2002b). Zhang et al. (2003) demonstrated that NLTCC antagonists inhibit morphine sensitization in mice and proposed that NLTCCs are involved in the development of morphine-induced neural and behavioral plasticity. In diabetic rats, nimodipine at 0.3–3.0 mg/kg i.p. potentiated the antinociceptve effects of morphine (Gullapalli et al., 2002). The route of administration and the duration of treatment appear to determine the ability of nimodipine to potentiate morphine. Lee and Yoburn (2000) found that nimodipine when administered s.c. to mice by a minipump at 100 mg/kg/day over 7 days, but not by single s.c. administration at 100 mg/kg, potentiated morphine-induced analgesia. This effect of nimodipine is not specific for opioids. Nociception induced by 5-HTP in mice was potentiated by nimodipine, nifedipine, or verapamil (Liang et al., 2004), and nimodipine was found to enhance the antihyperalgesic effects of diclofenac in formalin pain model in rats (Sukriti and Pandhi, 2004). In an attempt to explain the mechanism of interaction of calcium channel antagonists with morphine, Shimizu et al. (2004) pretreated mice with high doses (40–80 mg/kg i.p.) of diltiazem, nimodipine, or verapamil prior to morphine (4 mg/kg s.c.) and found that these drugs potentiate analgesic effects and increase serum levels of morphine.

FUNCTION

17

By acute administration to healthy volunteers, diltiazem, nimodipine, or verapamil did not enhance the analgesic effects of morphine (Hasegawa and Zacny, 1997). In patients with cancer pain, nimodipine did not enhance morphine-induced analgesia (Roca et al., 1996). Also, in patients undergoing colorectal surgery, neither oral nifedipine nor intravenous nimodipine increased the analgesic potency of morphine (Zarauza et al., 2000). In patients undergoing knee replacement surgery, oral nimodipine increased morphine consumption without enhancing its analgesic effect (Casey et al., 2006). There is currently no definitive explanation for the apparent discrepancy between animal and human studies in respect to the ability of NLTCC antagonists to potentiate the analgesic effects of morphine. To better understand the nature of the discrepancy, the optimal blood levels of NLTCCs antagonists required for the enhancement of morphine analgesia in animals should be determined, so that the same blood levels can be achieved in clinical studies. It is conceivable that sustained release formulations of NLTCC antagonists administered for at least a week would enhance the analgesic effects of morphine. 1.3.4

Epilepsy

Epileptogenic activity in neurons is thought to be activated by an inward Ca2þ current. After entering neurons at least partially through NLTCCs, calcium ions are thought to regulate various aspects of synaptic activity, including epileptogenesis (De Lorenzo, 1986). The specific molecular mechanisms involved in the epileptogenesis are poorly understood, and the role of various subunits and isoforms of NLTCCs in the epileptogenic activity is not yet known. It has been suggested that antagonists of T-type calcium channels are effective in treatment of absence seizures, while NLTCCs may control partial seizures (Kułak et al., 2004). The NLTCC activator BAY K 8644 has proconvulsant activity in animals. Most antagonists of NLTCCs have been shown to have anticonvulsant activity at least in some animal models of epilepsy. Clinical data are contradictory. Anticonvulsant activity of nifedipine and nimodipine has been demonstrated in small open studies, but controlled studies with either nifedipine (Larkin et al., 1992) or nimodipine (Larkin et al., 1991; Meyer et al., 1995) were disappointing. Nimodipine has also been used in the prevention of eclampsia, but appears to be less effective than magnesium sulfate (Belfort et al., 2003). 1.3.5

Drug and Ethanol Dependence

NLTCCs appear to play a role in drug addiction and alcohol dependence, but the effectiveness of the antagonists in the treatment of drug addiction remains controversial. Nimodipine, 5–20 mg/kg s.c., or isradipine, 1–3 mg/kg s.c., inhibited selfadministration of morphine or cocaine in drug-na€ıve mice (Kuzmin et al., 1992). At 20 mg/kg s.c., nimodipine decreased the sensitivity of rodents to the reinforcing effects of cocaine (Kuzmin et al., 1996). Cocaine-induced elevation of plasma catecholamines was prevented by nimodipine in squirrel monkeys (Trouve et al., 1990). Nitrendipine, flunarizine, or diltiazem protected rats from convulsions and death caused by a large dose of cocaine (60 mg/kg i.p.) (Trouve and Nahas, 1990).

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Withdrawal signs in morphine-dependent rats were effectively antagonized by NLTCCs (Ramkumar and El-Fakahany, 1988). Chronic exposure to cocaine facilitated the function of L-type calcium channels in pyramidal neurons of medial prefrontal cortex of rats (Nasif et al., 2005). It appears that antagonists of NLTCCs may have a place in the treatment of cocaine toxicity or of withdrawal symptoms, but in cocaine-dependent patients craving for cocaine was not affected by nimodipine (Rosse et al., 1994). Dihydropyridine-type antagonists of NLTCCs were found to interact with ethanol in animals, reversing seizures and other symptoms associated with ethanol withdrawal (Little et al., 1986; Littleton et al., 1990), blocking self-administration of 5% (but not higher concentrations) of ethanol (Smith et al., 1999), and blocking tolerance to the antinociceptive effects of ethanol during withdrawal (Gatch, 2006). Physical dependence on alcohol is associated with an increased number of dihydropyridine-sensitive calcium channels in the rat brain (Dolin et al., 1987). More recently, Katsura et al. (2005, 2006) demonstrated that physical dependence on ethanol in mice is accompanied by increased expression (and possibly function) of Cav1.2 and Cav1.3 isoforms and a2/d1 subunits of NLTCCs in mouse brain. Selective antagonists of channels containing these isoforms may reduce or abolish the dependence on ethanol and possibly on other habit-forming substances. Antagonists of NLTCCs, nimodipine, verapamil, or diltiazem, attenuated nicotineinduced locomotor sensitization and place preference in mice (Biala, 2003). The same drugs attenuated the antinociceptive effects of nicotine as well as cross-tolerance to the antinociceptive actions of nicotine and morphine (Biala and Weglinska, 2006). These findings suggest that a common mechanism involving L-type calcium channels may be responsible for the development of tolerance to the antinociceptive effects of morphine and nicotine. Chronic administration of nicotine leads to upregulation of Cav1.2, Cav1.3, Cav1.4, and a2d 1 subunits in mouse brain (Hayashida et al., 2005). Anxiogenic effects of nicotine in mice as well as the development of tolerance to this effect were attenuated by nimodipine, flunarizine, verapamil, or diltiazem (Biala and Budzynska, 2006). Attenuation of nicotine effects by NLTCCs antagonists may not, however, be mediated solely by calcium channels. It has been recently shown that nimodipine and nifedipine can also block nicotinic acetylcholine receptors (nAChRs) directly (Wheeler et al., 2006). 1.3.6

Hearing

NLTCCs and specifically Cav1.3 isoform are essential for synaptic transmission in cochlear inner hair cells and hair cell development in mice (Brandt et al., 2003, 2005; Nemzou et al., 2006). Cav1.3/ mice are deaf and show outer hair cell loss at the apical cochlea, while heterozygous (Cav1.3þ/) mice have increased hearing threshold for low-frequency sounds (Dou et al., 2004). These findings indicate the importance of the Cav1.3 subtype of NLTCCs for normal hearing. Antagonists of NLTCCs could, therefore, be expected to impair hearing. However, nimodipine improved hearing in patients with sudden hearing loss (Handrock, 1985; Theopold, 1985). In rats, it improved cochlear microphonics (Jastreboff and Brennan, 1988) and prevented

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neuronal degeneration in the cochlear nerve (Sekiya et al., 2002), but did not protect gerbils from noise-induced hearing loss (Boettcher et al., 1998). 1.3.7

Vision

Light-dependent Ca2þ influx into photoreceptors in the retina is controlled by Cav1.4 and Cav1.3 subtypes of L-type calcium channels. These channels are located presynaptically in retinal synapses and control neurotransmitter release, primarily glutamate, but also GABA, nitric oxide, and dopamine (Barnes and Kelly, 2002). Cav1.3 is consistently expressed in AII amacrine cells, retinal neurons that have a critical role in night vision (Habermann et al., 2003). Mutations of the gene that encodes retinal Cav1.4-type channels are linked to the night blindness type 2 (CSNB2) (Hoda et al., 2006). No effect of NLTCCs on night vision has been described in the literature, but nimodipine was found to improve visual field and color vision in patients with normal tension glaucoma (Piltz et al., 1998). At 90 mg/day, nimodipine in combination with aspirin, 100 mg/day, improved visual field and hearing dysfunction in a patient with Susac’s syndrome (Wildemann et al., 1996). The pathogenesis of this syndrome is unknown, but its symptomatology includes memory loss, impaired hearing, and vision loss and may conceivably involve NLTCCs. 1.3.8

Gene Transcription

The most important and critical function of NLTCCs is probably the coupling of neuronal activity to gene transcription. Nuclear transcription factors (i.e., pCREB and NFATc44) are activated by Ca2þ influx via postsynaptic L-type calcium channels (Bito et al., 1996; Dolmetsch et al., 2001). In hippocampal neurons, at low levels of stimulation nuclear pCREB signaling is preferentially mediated by the Cav1.3 subtype of NLTCCs (Zhang et al., 2006). The mechanism linking calcium channels to genes involves calcium channel-associated transcription regulator (CCAT). It binds to nuclear proteins and regulates the expression of endogenous genes controlling neuronal signaling and excitability (Gomez-Ospina et al., 2006). CCAT increases dendritic length and promotes contacts between neurons and extracellular matrix. 1.3.9

Cell Differentiation

Ca2þ influx through NLTCCs also affects expression of genes involved in cell proliferation, programmed cell death, and differentiation of neurons. According to D’Ascenzo et al. (2006), differentiation of neural stem/progenitor cells (NSCs) isolated from brain cortex of newborn mice depends on the Ca2þ influx through NLTCCs containing Cav1 isoforms. Immature GABAergic neurons are particularly sensitive to low Ca2þ levels, and Ca2þ influx through L- and T-type channels protects immature neurons from apoptosis (Pardo and Honegger, 1999). Also, in cerebellar Purkinje neurons, Ca2þ influx through L-type channels appears to be more important in the early rather than in the late stages of their development (Gruol et al., 2006).

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1.3.10

Interactions of NLTCCs with Neurotransmitter Systems

The function of NLTCCs should not be viewed in isolation from other neurotransmitter systems. Ca2þ entry through NLTCCs is capable of modulating signaling of most neurotransmitter receptors, and many neurotransmitters can modulate the function of NLTCCs. Dopaminergic signaling plays a key role in the physiology and pathology of the central nervous system. D1 receptor-mediated CREB phosphorylation depends on NLTCCs. D1 receptor stimulation in striatal neurons reverses the effects of NLTCC antagonists on CREB phosphorylation, so that antagonists promote rather than block Ca2þ entry into the neurons (Eaton et al., 2004). Interdependence between dopaminergic and NMDA receptors and NLTCCs has been demonstrated (Cepeda and Levine, 1998). Cannabinoid receptor CB1 was described to inhibit calcium influx through NLTCCs in neonatal rat nucleus solitarius (Endoh, 2006), and opioids were found to modulate NLTCCs through orphan opioid receptor activation (Hurl
e et al., 1999). The extent of the interplay between receptors and neurotransmitters is not yet fully appreciated and the molecular mechanisms of these interactions are not yet completely understood, but modulation of other neurotransmitter systems is likely to be an important function of not only NLTCCs but also all ion channels.

1.4 CHANNELOPATHIES AND INHERITED DISORDERS Over the past two decades, an increasing number of mutations in voltage-gated calcium channels have been mapped and linked to a number of inherited disorders. A number of comprehensive reviews are available (inter alia, Lorenzo and Beam, 2000, 2005; Muth et al., 2001; Pietrobon, 2002; Striessnig et al., 2004; Pietrobon, 2005; Biduad et al., 2006; Bracey and Wray, 2006; Cannon, 2006). However, the majority of these mutations and inherited disorders are associated with non-L-type channel genes (Cav2 and Cav3), L-type channels, or associated proteins involved in muscle (skeletal, smooth, and cardiac) function. Associated with Cav2 and Cav3 genes are hemiplegic migraine, several ataxias, juvenile myocolonic epilepsy, and idiopathic generalized epilepsies, including childhood absence epilepsy. Associated with skeletal muscle are (1) hypokalemic periodic paralysis (hypoPP), a periodic muscle weakness associated with reduced serum Kþ levels and linked to missense mutations in Cav1.1 (a1S); (2) malignant hyperthermia (MH) linked to multiple mutations in the ryanodine receptor associated with the voltage-sensing dihydropyridine receptor and linked to life-threatening body temperature increases during a number of pharmacological interventions, including general anesthesia and skeletal muscle relaxants; and (3) central core disease, also associated with defects in ryanodine receptors. Two cardiac muscle disorders, arrhythmogenic right ventricular cardiomyopathy and familial polymorphic ventricular tachycardia, are also linked to defects in the ryanodine receptor. Defects in L-type channels associated with neuronal disorders have been described for Cav1.2 and particularly for Cav1.4. A mutation in the Cav1.2 gene is associated with childhood disorder termed “Timothy syndrome’’ associated with multiple

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electrophysiological defects and sudden death caused by cardiac arrhythmias and contributed by gain of function mutations with reduced channel inactivation (Splawski et al., 2004, 2005) and aberrant phosphorylation of Ser49 in the S6 helix of domain 1 (Erxleben et al., 2006). These individuals have webbed fingers and toes (syndactyly), cognitive abnormalities, and autism. The latter pathology is of particular interest although the generalized defect produced by this mutation makes attribution to a specific neuronal pathway difficult. By contrast, mutations in the Cav.4 channel, almost totally distributed in and linked to retinal function (Bech-Hansen et al., 1998; Strom et al., 1998), have been well described as linked to incomplete congenital stationary night blindness (CSNB2) and characterized by varying levels of night blindness and a reduced visual acuity. Some 60 mutations have been described: approximately 50% are missense mutations leading to nonfunctional proteins, while the remainder lead to expressed channels with varying levels of dysfunction (McRory et al., 2004; Hoda et al., 2006). It is likely that more mutations will be discovered in Cav1 channels linked to neuronal dysfunction, although difficulties exist because this class of channels is so widely expressed (Gargus, 2006). However, knockout studies will provide valuable leads (Muth et al., 2001). Thus, mice lacking Cav1.3 channels have both sinoatrial node dysfunction and congenital deafness, the latter being associated with degeneration of hair cells (Platzer et al., 2000; Nemzou et al., 2006). Mice lacking the Cav1.2 gene have impaired glucose tolerance and insulin secretion, the latter reflecting the absence of an exocytotic component of fusion of secretory granule attached to this channel (Schulla et al., 2003). In addition, it is highly plausible that since the multiple cellular calcium signaling and regulatory mechanisms are closely linked, changes in one may lead to compensating changes in others. Thus, a missense mutation in the Cav2.1 channel underlies the behavior of tottering mice that have ataxia, paroxysmal dystonia, and spontaneous behavioral arrest (Pietrobon, 2002). Since the inducible dystonia component of this mutation can be blocked by L-type antagonists, including diltiazem, nifedipine, and verapamil, and since Cav1.2 channels are significantly upregulated in Purkinje and cerebellar cells, it appears that Cav2.1 dysfunction has produced a compensating upregulation of Cav1.2 channels (Campbell and Hess, 1999). Similarly, Cav1.3 knockout mice have impaired pancreatic islet cell function but have a compensatory overexpression of the Cav1.2 gene (Namkung et al., 2001).

1.5 CLINICAL IMPLICATIONS AND FUTURE PERSPECTIVES In spite of important advances in the physiology and molecular biology of calcium channels during the past 20 years, very little progress has been made in the development of calcium channel ligands as drugs for the treatment of central nervous system diseases. No clinical use for the available activators of NLTCCs has been found. BAY K 8644 and similar activators of NLTCCs produce dystonia, self-injurious behavior, and convulsions in rodents (Jinnah et al., 2000; Kasim and Jinnah, 2003) and are unlikely to be tried in humans.

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The antagonists, while widely used in cardiovascular diseases, have found only limited use in the treatment of CNS diseases (Scriabine et al., 1989, 1991; Scriabine, 2002). As noted in a recent review (Triggle, 2007), in the CNS a variety of calcium channel types contribute to neuronal activity and brain damage is likely to involve multiple pathways, so that blockade of a single channel type may not be the right strategy. Also, the failure of nimodipine in pivotal trials in either Alzheimer’s disease or stroke (Mohr, 1991; Morich et al., 1996) discouraged further trials with antagonists of NLTCCs in these conditions. Subarachnoidal hemorrhage is currently the only FDA-approved indication for nimodipine in the United States. This drug is, however, prescribed “off-label’’ in the prevention of migraine attacks and cluster headache in the United States and is approved for the treatment of “organic brain syndrome’’ in Germany and some of the other European countries. Further clinical trials with NLTCC antagonists in migraine as well as in drug and ethanol addiction should be considered. Such studies will probably be conducted with novel compounds, selective for subunits or isoforms of NLTCCs, or compounds with multiple sites of action. There are no selective NLTCCs yet, but Kiewert et al. (2006) described the pharmacology of NGP1-01, a polycyclic amine, which blocks NLTCCs as well as NMDA channels. Its neuroprotective potency is similar to that of nimodipine and it is structurally related to memantine, so the authors suggested its possible usefulness in stroke as well as in the treatment of neurodegenerative diseases. There is an obvious need for more basic and translational research on the function of NLTCC isoforms and their ligands. The observed interactions of NLTCCs with neurotransmitters or their receptors suggest the use of NLTCCs as adjuncts in the therapy of CNS diseases.

1.6 SUMMARY L-type calcium channels are widely distributed in the central nervous system. Like in other tissues, the channels consist of the pore-forming a1 and three regulatory subunits (a2, b, and g). Three isoforms of a1 subunit have been identified in the central nervous system: Cav1.2, Cav1.3, and Cav1.4. Channels with different isoforms of the a1 subunit differ in some of their functions. Cav1.2-containing channels appear to be involved in cognition and memory. Cav1.3-containing channels have many functions similar to those of Cav1.2-containing channels, but are essential for the synaptic transmission in cochlear inner hair cells and control hearing. Cav1.4-containing channels control Ca2þ influx into photoreceptors of the retina and are involved in the control of vision. L-type calcium channels are also involved in pain perception, neuronal excitability, gene transcription, and cell differentiation. They interact with other transmitter and receptor systems. Activators of L-type calcium channels are proconvulsant and neurotoxic. Antagonists are neuroprotective but have thus far found only limited use in the treatment of CNS diseases. In the United States, only nimodipine has been approved for the prevention of neurological deficits following subarachnoidal hemorrhage. L-type calcium channel antagonists, including nimodipine, appear to be effective in

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the prevention of migraine and cluster headache and can conceivably find new applications in the treatment of dementia, in drug and alcohol dependence, or as adjuncts in the treatment of epilepsy. Specific inhibitors of channels with different isoforms should be developed. Cav1.3a1 antagonists can be expected to have antidepressant and/or anxiolytic properties.

REFERENCES Barnes S, Kelly MEM, 2002. Calcium channels at the photoreceptor synapse. Adv Exp Med Biol 514: 465–476. Baumann L, Gerstner A, Zong X, Biel M, Wahl-Schott C, 2004. Functional characterization of the L-type Ca2þ channel Cav1.4a1 from mouse retina. Invest Ophthalmol Vis Sci 45: 708–713. Bean BP, 1984. Nitrendipine block of cardiac calcium channels. High affinity binding to the inactivated state. Proc Natl Acad Sci USA 81: 6388–6392. Bean BP, 1989. Classes of Ca2þ channels in vertebrate cells. Annu Rev Physiol 51: 367–384. Bech-Hansen NT, Naylor MJ, Maybaum TA, Pearce WG, Koop B, Fishman GA, Mets M, Musarella MA, Boycott KM, 1998. Loss-of-function mutations in a calcium-channel alpha1-subunit gene in Xp11.23 cause incomplete X-linked congenital stationary night blindness. Nat Genet 19: 264–267. Belfort MA, Anthony J, Saade GR, Allen JC Jr, 2003. A comparison of magnesium sulfate and nimodipine for the prevention of eclampsia. N Engl J Med 348: 304–311. Bi G-Q, Poo M-M, 1998. Synaptic modifications in cultured hippocampal neurons: dependence on spike timing, synaptic strength and postsynaptic cell type. J Neurosci 18: 10464– 10472. Biala G, 2003. Calcium channel antagonists suppress nicotine-induced place preference and locomotor sensitization in rodents. Pol J Pharmacol 55: 327–335. Biala G, Budzynska B, 2006. Reinstatement of nicotine-conditioned place preference by drug priming: effects of calcium channel antagonists. Eur J Pharmacol 537: 85–93. Biala G, Weglinska B, 2006. On the mechanism of cross-tolerance between morphine and nicotine-induced nociception: involvement of calcium channels. Prog Neuropsychopharmacol Biol Psychiatry 30: 15–21. Biduad I, Mezghrani A, Swayne LA, Monteil A, Lory P, 2006. Voltage-gated calcium channels in genetic diseases. Biochim Biophys Acta 1763: 1169–1174. Bito H, Deisseroth K, Tsien RW, 1996. CREB phosphorylation and dephosphorylation: a Ca2þand stimulus duration-dependent switch for hippocampal gene expression. Cell 67: 1203– 1214. Boettcher FA, Caldwell RK, Gratton MA, White DR, Miles LR, 1998. Effects of nimodipine on noise-induced hearing loss. Hear Res 121: 139–146. Bourient E, Mangoni ME, Nargeot J, 2004. Dissecting the functional role of different isoforms of the L-type Ca2þ channel. J Clin Invest 113: 1382–1384. Bracey K, Wray D, 2006. Genetic and acquired channelopathies. In: Triggle, DJ, Gopalakrishnan M, Rampe D, Zheng W (Eds), Voltage-Gated Ion Channels as Drug Targets, WileyVCH, Weinheim, Germany, pp. 381–427.

24

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

Brandt A, Striessnig J, Moser T, 2003. Cav1.3 channels are essential for development and presynaptic activity of cochlear inner hair cells. J Neurosci 23: 10832–10840. Brandt A, Khimich D, Moser T, 2005. Few Cav1.3 channels regulate the exocytosis of a synaptic vesicle at the hair cell ribbon synapse. J Neurosci 25: 11577–11585. Brosenitsch TA, Salgado-Commissariat D, Kunze DL, Katz DM, 1998. A role for L-type calcium channels in developmental regulation of transmitter phenotype in primary sensory neurons. J Neurosci 18: 1047–1055. Budriesi R, Cosimelli B, Ioan P, Carosati E, Ugenti MP, Spisani R, 2007. Diltiazem analogues: the last ten years on structure–activity relationships. Curr Med Chem 14: 279–287. Campbell DB, Hess EJ, 1999. L-type calcium channels contribute to the tottering mouse dystonic episodes. Mol Pharmacol 55: 23–31. Cannon SC, 2006. Pathomechanisms in channelopathies of skeletal muscle and brain. Annu Rev Neurosci 29: 387–415. Cao Y-Q, 2006. Voltage-gated calcium channels and pain. Pain 126: 5–9 Casey G, Nortcliffe SA, Sharpe P, Buggy DJ, 2006. Perioperative nimodipine and postoperative analgesia. Anesth Analg 102: 504–508. Catterall WA, 2000. Structure and regulation of voltage-gated calcium channels. Annu Rev Cell Biol 16: 521–555. Catterall WA, 2005. Biochemical studies of voltage-gated Ca2þ channels. In: Zamponi G (Ed.), The Voltage-Gated Calcium Channel, Kluwer Academic/Plenum Publishing, New York, pp. 48–60. Catterall WA, Perez-Reyes E, Snutch TP, Striessnig T, 2005. International Union of Pharmacology. XLVIII, Nomenclature and structure–function relationships of voltage-gated calcium channels. Pharmacol Rev 57: 411–425. Cens T, Restitutio S, Rousset M, Charnet P, 2005. Role of b subunits in voltage-gated calcium channel function. In: Zamponi G (Ed.), Voltage-Gated Calcium Channels, Kluwer Academic/Plenum Publishing, New York, pp. 95–112. Cepeda C, Levine MS, 1998. Dopamine and N-methyl-D-aspartate receptor interaction in the neostriatum. Dev Neurosci 20: 1–18. Cifuentes F, Licona II, DeLeon L, Medina P, De-Miguel FF, Morales MA, 2004. Contribution of different calcium channels to long-term potentiation in superior cervical ganglion of the rat. Neuroscience 129: 647–653. Clark NC, Nagano N, Kuenzi FM, Jarolimek W, Huber I, Walter D, Wietzorrek G, Boyce S, Kullmann DM, Striessnig J, et al., 2003. Neurological phenotype and synaptic function in mice lacking Cav1.3 alpha subunit of neuronal L-type voltage-dependent Ca2þ channels. Neuroscience 120: 435–442. Cohen CJ, Spires S, van Skiver D, 1992. Block of T-type channels in guinea-pig atrial cells by antiarrhythmic agents and calcium antagonists. J Gen Physiol 100: 703–728. D’Ascenzo M, Piacentini R, Casalbore P, Budoni M, Pallini R, Azzena GB, Grassi C, 2006. Role of L-type Ca2þ channels in neural stem/progenitor cell differentiation. Eur J Neurosci 23: 935–944. De Jongh KS, Warner C, Catterall WA, 1990. Subunits of purified calcium channels. Alpha2 and delta are encoded by the same gene. J Biol Chem 265: 14738–14741. De Lorenzo RJ, 1986. A molecular approach to the calcium signal in brain: relationship to synaptic modulation and seizure discharge. Adv Neurol 44: 435–464.

REFERENCES

25

Del Pozo E, Caro G, Baeyens JM, 1987. Analgesic effects of several calcium channel blockers in mice. Eur J Pharmacol 137: 155–160. Deisseroth K, Mermelstein PG, Xia H, Tsien RW, 2003. Signaling from synapse to nucleus: the logic behind the mechanisms. Curr Opin Neurobiol 13: 34–365. Disterhoft JF, Moyer JR, Thompson LT, 1994. The calcium rationale in aging and Alzheimer’s disease. Evidence from an animal model of normal aging. Ann NY Acad Sci 747: 382– 406. Doering C, Zamponi G, 2006. Calcium channels: overview of voltage-gated calcium channels. In: Triggle DJ, Gopalakrishnan M, Rampe D, Zheng W (Eds), Voltage-Gated Ion Channels as Drug Targets, Wiley-VCH, Mannheim, pp. 65–83. Dolin SJ, Little HJ, Hudspith M, Pagonis C, Littleton JM, 1987. Increased dihydropyridine sensitive calcium channels in rat brain may underlie ethanol physical dependence. Neuropharmacology 26: 275–279. Dolmetsch RE, Pajvani U, Fife K, Spotts JM, Greenberg ME, 2001. Signaling to the nucleus by an L-type calcium channel–calmodulin complex through the MAP kinase pathway. Science 284: 333–339. Dou H, Vazquez AE, Namkung Y, Chu H, Cardell EL, Nie L, Parson S, Shin H-S, Yamoah EN, 2004. Null mutation of alpha1D Ca2þ channel gene results in deafness but no vestibilar defect in mice. JARO 5: 215–226. Eaton ME, Macias W, Youngs RM, Rajadhyaksha A, Dudman JT, Konradi C, 2004. L-type Ca2þ channel blockers promote Ca2þ accumulation when dopamine receptors are activated in striatal neurons. Brain Res Mol Brain Res 131: 65–72. Endoh T, 2006. Pharmacological characterization of inhibitory effects of postsynaptic opioid and cannabinoid receptors on calcium currents in neonatal rat nucleus tractus solitarius. Br J Pharmacol 147: 391–401. Erxleben C, Liao Y, Genetile S, Chin D, Gomez-Alegria C, Mori Y, Birnbaumer L, Armstrong D, 2006. Cyclosporin and Timothy syndrome increase mode 2 gating of CaV1.2 calcium channels through aberrant phosphorylation of S6 helices. Proc Natl Acad Sci USA 103: 3932–3937. Evans RM, Zamponi GW, 2006. Presynaptic Ca2þ channels—integration centers for neuronal signaling pathways. Trends Neurosci 29: 617–623. Facchinetti F, Fasolato C, Del Giudice E, Burgo A, Furegato S, Fusco M, Basso E, Seraglia R, D’Arrigo A, Leon A, 2006. Nimodipine selectively stimulates b-amyloid1–42 secretion by a mechanism independent of calcium influx blockade. Neurobiol Aging 27: 218–227. Field MJ, Cox PJ, Stott E, Melrose H, Offord J, Su TZ, Bramwell S, Corradini L, England S, Winks J, et al., 2006. Identification of the a2d1 subunit of voltage-dependent calcium channels as a molecular target for pain mediating the analgesic action of pregabalin. Proc Natl Acad Sci USA 103: 17537–17542. Fischhof PK, 1993. Divergent neuroprotective effects of nimodipine in PDD and MID provide indirect evidence of disturbance in Ca2þ homeostasis in dementia. Methods Find Exp Clin Pharmacol 15: 549–555. Fleckenstein A, 1983. Calcium Antagonists in Heart and Smooth Muscle, Wiley, New York. Forette F, Seux M-L, Staessen JA, Thijs L, Birkenhager WH, Babarskleine MR, Babeanu S, Bossine A, Gil-Extremera B, Girerd X, et al., 1998. Prevention of dementia in randomised double-blind placebo controlled systolic hypertension in Europe (Syst-Eur) trial. Lancet 352: 1347–1351.

26

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

Frampton JE, Foster RH, 2005. Pregabalin in the treatment of postherpetic neuralgia. Drugs 65: 111–118. Gargus JJ, 2006. Ion channel functional candidate genes in multigenic neuropsychiatric disease. Biol Psychiatry 60: 77–185. Gatch MB, 2006. Tolerance to the antinociceptive effects of ethanol during ethanol withdrawal. Prog Neuropsychopharmacol Biol Psychiatry 30: 946–952. Goldmann S, Stoltefuss J, 1991. 1,4-Dihydropyridines: effects of chirality and conformation on the calcium antagonist and calcium agonist activities. Angew Chem Int Ed 30: 1559–1578. Gomez-Ospina N, Tsuruta F, Barreto-Chang O, Hu L, Dolmetsch R, 2006. The C-terminus of the L-type voltage-gated calcium channel Cav1.2 encodes a transcription factor. Cell 127: 591–606. Gruol DL, Netzeband JG, Schneeloch J, Gullette CE, 2006. L-type Ca2þ channels contribute to current-evoked spike firing and associated Ca2þ signals in cerebellar Purkinje neurons. Cerebellum 5: 146–154. Gullapalli S, Ramarao P, 2002a. Regulation of dihydropyridine-sensitive Ca2þ channels during naloxone-induced opioid supersensitivity in rats. Eur J Pharmacol 451: 271–277. Gullapalli S, Ramarao P, 2002b. L-type Ca2þ channel modulation by dihydropyridines potentiates k-opioid receptor agonist-induced acute analgesia and inhibits development of tolerance in rats. Neuropharmacology 42: 467–475. Gullapalli S, Gurumoorthy K, Kaul CL, Ramarao P, 2002. Role of L-type Ca2þ channels in attenuated morphine antinociception in streptozotocin-diabetic rats. Eur J Pharmacol 435: 187–194. Habermann CH, O’Brien J, W€assle H, Protti DA, 2003. AII amacrine cells express L-type calcium channels at their output synapses. J Neurosci 36: 6904–6913. Handrock M, 1985. Sudden loss of hearing. In: Betz E, Deck K, Hoffmeister F (Eds), Nimodipine: Pharmacological and Clinical Properties, Schattauer Verlag, Stuttgartt, pp. 481–496. Hanson JE, Smith Y, 2002. Subcellular distribution of high-voltage activated calcium channel subtypes in rat globus pallidus neurons. J Comp Neurol 442: 89–98. Hasegawa AE, Zacny JP, 1997. The influence of three L-type calcium channel blockers on morphine effects in healthy volunteers. Anesth Analg 85: 633–638. Hayashida S, Katsura M, Torigoe F, Tsujimura A, Ohkuma S, 2005. Increased expression of Ltype high voltage-gated calcium channel alpha1 and alpha2/delta subunits in mouse brain after chronic nicotine administration. Brain Res Mol Brain Res 135: 280–284. Hell JW, Westenbroek RE, Warner C, Ahlijanian MK, Prystay W, Gilbert MM, Snurch TP, Catterall WA, 1993. Identification and subcellular localization of the neuronal class C and class D L-type calcium channel a1 subunits. J Cell Biol 123: 949–962. Helton TD, Xu W, Lipscombe D, 2005. Neuronal L-type calcium channels open quickly and are inhibited slowly. J Neurosci 25: 10247–10251. Herlitze S, Mark MD, 2005. Distribution and targeting mechanisms of voltage activated Ca2þ channels. In: Zamponi G (Ed.), Voltage-Gated Calcium Channels, Kluwer Academic/ Plenum Press, New York, pp. 112–140. Hess P, 1990. Ca2þ channels in vertebrate cells. Annu Rev Neuroscience 13: 337–356. Hille B, 1977. Local anesthetics: hydrophilic and hydrophobic pathways for the drug–receptor interaction. J Gen Physiol 69: 497–515.

REFERENCES

27

Hoda J-C, Zaghetto F, Singh A, Koschak A, Striessnig J, 2006. Effects of congenital stationary night blindness type 2 mutations R508Q and L1364H on Cav1.4 L-type Ca2þ channel function and expression. J Neurochem 96: 1648–1658. Hoffmeister F, Tettenborn D, 1986. Calcium agonists and antagonists of the dihydropyridine type: antinociceptive effects, interference with opiate m receptor agonists and neuropharmacological actions in rodents. Psychoneuropharmacology 90: 299–307. Hofmann F, Lacinova L, Klugbauer N, 1999. Voltage-dependent calcium channels: from structure to function. Rev Physiol Biochem Pharmacol 139: 33–87. Hondeghem LM, Katzung BG, 1985. Anti-arrhythmic agents: the modulated receptor mechanism of action of sodium and calcium channel blocking drugs. Annu Rev Pharmacol Toxicol 24: 387–423. Hu H, Marban E, 1998. Isoform-specific inhibition of L-type calcium channels by dihydropyridines is independent of isoform-specific gating properties. Mol Pharmacol 53: 902–907. Hurl
e MA, S
anchez A, Garc
ıa-Sancho J, 1999. Effects of k- and m-opioid receptor agonists on Ca2þ channels in neuroblastoma cells: involvement of the orphan opioid receptor. Eur J Pharmacol 379: 191–198. Janis RA, Triggle DJ, 1983. New developments in Ca2þ antagonists. J Med Chem 26: 775–785. Janis RA, Triggle DJ, 1984a 1,4-Dihydropyridine Ca2þ channel antagonists and activators. A comparison of binding characteristics with pharmacology. Drug Dev Res 4: 257–285. Janis RA, Triggle DJ, 1984b Calcium channel antagonists: new perspectives from the radioligand binding assay. In: Back N, Spector, S (Eds), Modern Methods in Pharmacology, Vol. 2. Liss, New York, pp. 1–28. Janis RA, Silver P, Triggle DJ, 1987. Drug action and cellular calcium regulation. Adv Drug Res 16: 309–591. Jastreboff PJ, Brennan JF, 1988. Specific effects of nimodipine on the auditory system. Ann NY Acad Sci 522: 716–718. Jinnah HA, Sepkuty JP, Ho T, Yitta S, Drew T, Rothstein JD, Hess EJ, 2000. Calcium channel agonists and dystonia in the mouse. Mov Disord 15: 542–551. Joshi I, Taylor CP, 2006. Pregabalin action at a model synapse: binding to presynaptic calcium channel a2-d subunit reduces neurotransmission in mice. Eur J Pharmacol 553: 82–88. Kandel ER, 2001. The molecular biology of memory storage: a dialogue between genes and synapses. Science 294: 1030–1038. Kasim S, Jinnah HA, 2003. Self-biting induced by activation of L-type calcium channels in mice: dopaminergic influences. Dev Neurosci 25: 20–25. Katsura M, Torigoe F, Hayashida S, Honda T, Tsujimura A, Ohkuma S, 2005. Ethanol physical dependence is accompanied by up-regulated expression of L-type high voltage calcium channel a1 subunits in mouse brain. Brain Res 1039: 211–215. Katsura M, Shibasaki M, Hayashida S, Torigoe F, Tsujimura A, Ohkuma S, 2006. Increase in expression of a1 and a2/d1 subunits of L-type high voltage-gated calcium channels after sustained ethanol exposure in cerebral cortical neurons. J Pharmacol Sci 102: 221–230. Khachaturian ZS, 1995. Calcium hypothesis of Alzheimer’s disease and brain aging. Ann NY Acad Sci 747: 1–11. Kiewert C, Hartmann J, Stoll J, Thekkumkara TJ, Van der Schyf CJ, Klein J, 2006. NGP-01 is a brain-permeable dual blocker of neuronal voltage- and ligand-operated calcium channels. Neurochem Res 31: 395–399.

28

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

Kordasiewicz HB, Thompson RM, Clark HB, Gomez CP, 2006. C-termini of P/Q-type Ca2þ channel alpha1a subunits translocate to nuclei and promote polyglutamine-mediated toxicity. Hum Mol Genet 15: 1587–1599. Kułak W, Sobaniec W, Woital K, Czuczwar J, 2004. Calcium modulation in epilepsy. Pol J Pharmacol 56: 29–41. Kumar PH, Stotz SC, Paramashivappa R, Beedle AM, Zamponi GW, Rao AS, 2002. Synthesis and evaluation of a new class of nifedipine analogs with T-type calcium channel binding activity. Mol Pharmacol 61: 649–658. Kuzmin A, Zvartau E, Gessa GL, Martellotta MC, Fratta W, 1992. Calcium antagonists isradipine and nimodipine suppress cocaine and morphine self-administration in drug-naive mice. Pharmacol Biochem Behav 41: 497–500. Kuzmin A, Semenova S, Ramsey NF, Zvartau EE, Van Ree JM, 1996. Modulation of cocaine intravenous self-administration in drug-na€ıve animals by dihydropyridine Ca2þ channel modulators. Eur J Pharmacol 295: 19–25 Lacinova L, 2005. Voltage-dependent calcium channels. Gen Physiol Biophys 24 (Suppl 1): 1–78. Lacinova L, Klugbauer N, Hofmann F, 2000. State- and isoform-independent interaction of isradipine with the a1C L-type calcium channel. Pfl€ ugers Arch 440: 50–60. Lai HC, Jan LY, 2006. The distribution and targeting of neuronal voltage-gated ion channels. Nat Rev Neurosci 7: 548–562. Landfield PW, 1987. Increased hippocampal Ca2þ channel activity in brain aging and dementia. Ann NY Acad Sci 747: 351–364. Larkin JG, McKee PJ,W, Blacklaw J, Thomson CG, Morgan IC, Brodie MJ, 1991. Nimodipine in refractory epilepsy: a placebo-controlled, add-on study. Epilepsy Res 9: 71–77. Larkin JG, Besag FM, Cox A, Williams J, Brodie MJ, 1992. Nifedipine for epilepsy? A doubleblind, placebo-controlled trial. Epilepsia 33: 2346–2352. Lee A, Catterall WA, 2005. Ca2þ-dependent modulation of voltage-gated Ca2þ channels. In: Zamponi G (Ed.), Voltage-Gated Calcium Channels, Kluwer Academic/Plenum Publishers, New York, pp. 183–193. Lee SC, Yoburn BC, 2000. The effect of nimodipine on opioid antagonist-induced upregulation and supersensitivity. Pharmacol Biochem Behav 66: 347–351. LeVere TE, Sandin M, 1989. Nimodipine in animal models of learning and memory. In: Traber J Gispen WH (Eds), Nimodipine and Central Nervous System Function: New Vistas. Schattauer Verlag, Stuttgart, pp. 163–174. Liang J-H, Li J-X, Wang X-H, Chen B, Lu Y, Zhang P, Han R, Ye X-F, 2004. L-type calcium channel blockers enhance 5-HTP-induced antinociception in mice. Acta Pharmacol Sin 25: 644–650. Lin C-H, Yeh S-H, Leu T-H, Chang ST, Wang P-W, 2003. Identification of calcineurin as a key signal in the extinction of fear memory. J Neurosci 23: 1574–1579. Lipscombe D, 2002. L-type calcium channels. Highs and lows. Circ Res 90: 933–935. Lipscombe D, Helton TD, Xu W, 2004. L-type calcium channels: the low down. J Neurophysiol 92: 2633–2641. Little HJ, Dolin SJ, Halsey MJ, 1986. Calcium channel antagonists decrease the ethanol withdrawal syndrome. Life Sci 39: 2059–2065.

REFERENCES

29

Littleton JM, Little HJ, Whittington MA, 1990. Effects of dihydropyridine calcium antagonists in ethanol withdrawal: doses required, stereospecificity and actions of Bay K 8644. Psychopharmacology 100: 387–392. Lorenzo NM, Beam KG, 2000. Calcium channelopathies. Kidney Int 57: 794–802. Lorenzo N, Beam KG, 2005. Calcium channelopathies. In: Zamponi G (Ed.), Voltage-Gated Calcium Channels, Kluwer Academic/Plenum Publishers, New York, pp. 240–261. Ludwig A, Flockerzi V, Hofmann F, 1997. Regional expression and subcellular localization of the a1 and b subunit of high voltage-activated calcium channels in rat brain. J Neurosci 17: 1339–1349. Lynch JJ III, Honore P, Anderson DJ, Bunelle WH, Mortell KH, Zhong C, Wadfe CL, Zhu CZ, Xu H, Marsh KC, et al., 2006. (L)-Phenylglycine, but not necessarily other a2d subunit voltage-gated channel ligands, attenuates neuropathic pain in rats. Pain 125: 136–142. Mansuy IM, 2003. Calcineurin in memory and bidirectional plasticity. Biochem Biophys Res Commun 311: 1195–1208. Mansuy IM, Mayford M, Jacob B, Kandel ER, Bach M, 1998. Restricted and regulated overexpression reveals calcineurin as a key component of the transition from short-term to long-term memory. Cell 92: 257–265. Martin MI, Del Val VL, Colado MI, Goicoechea C, Alfaro MJ, 1996. Behavioral and analgesic effects induced by administration of nifedipine and nimodipine. Pharmacol Biochem Behav 55: 93–98. McDonough S, 2004. Calcium Channel Pharmacology. Kluwer Academic/Plenum Publishers, New York. McDonough S, Bean BP, 2006. State-dependent drug interactions with ion channels. In: Triggle DJ, Gopalakrishnan M, Rampe D, Zheng W (Eds), Voltage-Gated Ion Channels as Drug Targets. Wiley-VCH, Weinheim, Germany, pp. 19–36. McRory JE, Hamid J, Doering CJ, Garcia E, Parker R, Hamming K, Chen L, Hildebrand M, Beedle AM, Feldcamp L, 2004. The CACNA1F gene encodes an L-type calcium channel with unique biophysical properties and tissue distribution. J Neurosci 24: 1707–1718. Meyer FB, Cascino GD, Whisnant JP, Sharbrough FW, Ivnik RJ, Gorman DA, Windschitl WL, So EL, O’Fallon WM, 1995. Nimodipine as an add-on therapy for intractable epilepsy. Mayo Clin Proc 70: 623–627. Miljanich GP, Ramachandran J, 1995. Antagonists of neuronal Ca2þ channels: structure, function and therapeutic implications. Annu Rev Pharmacol Toxicol 35: 707–734. Miranda HF, Bustamante D, Kramer V, Pelissier T, Saavedra H, Paelle C, Fernandez E, Pinardi G, 1992. Antinociceptive effects of Ca2þ channel blockers. Eur J Pharmacol 217: 137–141. Miranda HF, Pelissier T, Sierralta F, 1993. Analgesic effect of intracerebroventricular administration of calcium channel blockers in mice. Gen Pharmacol 24: 201–204. Mohr JP, 1991. Nimodipine and other calcium antagonists in acute ischemic stroke. In: Scriabine FA, Teasdale GM, Tettenborn D, Young W (Eds), Nimodipine: Pharmacological and Clinical Results in Cerebral Ischemia. Springer Verlag, Heidelberg, pp. 151–164. Moosmang S, Haider N, Klugbauer N, Adelsberger H, Langwieser N, M€ uller J, Stiess M, Marais E, Schulla V, Lacinova L, et al., 2005a. Role of hippocampal Cav1.2 Ca2þ channels in NMDA receptor-independent synaptic plasticity and spatial memory. J Neurosci 25: 9883–9892.

30

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

Moosmang S, Lenhardt P, Haider N, Hofmann F, Wegener JW, 2005b. Mouse models to study Ltype calcium channel function. Pharmacol Ther 106: 347–355. Morel N, Buryi V, Feron O, Gomez J-P, Christen M-O, Godfraind T, 1998. The action of calcium channel blockers on recombinant L-type calcium channel a1-subunits. Br J Pharmacol 125: 1005–1012. Morich FJ, Bieber F, Lewis JM, et al., 1996. Nimodipine in treatment of probable Alzheimer’s disease. Clin Drug Invest 11: 185–195. Muth JN, Varadi G, Schwartz A, 2001. Use of transgenic mice to study voltage-dependent Ca2þ channels. Trends Pharmacol Sci 22: 526–531. Namkung Y, Skrypnyk N, Jeong M-J, Lee T, Lee M-S, Kim H-L, Chin H, Suh P-G, Kim S-S, Shin H-S, 2001. Requirement for the L-type Ca2þ channel a1D subunit in postnatal pancreatic b-cell generation. J Clin Invest 108: 1015–1022. Nasif FJ, Hu X-T, White FJ, 2005. Repeated cocaine administration increases voltage-sensitive calcium currents in response to membrane depolarization in medial prefrontal cortex pyramidal neurons. J Neurosci 25: 3674–3679. Nemzou NRM, Bulankina AV, Khimich D, Giese A, Moser T, 2006. Synaptic organization in cochlear inner hair cells deficient for the Cav1.3 (alpha1D) subunit of L-type Ca2þ channels. Neuroscience 141: 1849–1860. Olson P, Tkatch T, Hernandez-Lopez S, Ulrich S, Ilijic E, Mugnaini E, Zhang H, Bezprozvanny I, Surmeier DJ, 2005. G-protein-coupled receptor modulation of striatal Cav1.3 Ca2þ channels is dependent on a Shank-binding domain. J Neurosci 25: 1050–1062. Pardo B, Honegger P, 1999. High sensitivity of immature GABAergic neurons to blockers of voltage-gated calcium channels. Dev Brain Res 115: 9–16. Pierrot N, Ghisdal P, Caumont A-S, Octave J-N, 2004. Intraneuronal Ab1–42 production triggered by sustained increase in cytosolic calcium concentration induces neuronal death. J Neurochem 88: 1140–1150. Pietrobon D, 2002. Calcium channels and channelopathies of the central nervous system. Mol Neurobiol 13: 254–265. Pietrobon D, 2005. Function and dysfunction of synaptic calcium channels: insights from mouse models. Curr Opin Neurobiol 15: 257–265. Piltz JR, Bose S, Lanchoney D, 1998. The effect of nimodipine, a centrally acting calcium antagonist, on visual function and muscular blood flow in patients with normal-tension glaucoma. J Glaucoma 7: 336–342. Pinard CR, Mascagni F, McDonald AJ, 2005. Neuronal localization of Cav1.2 L-type calcium channels in rat basolateral amygdala. Brain Res 1064: 52–55. Platzer J, Engel J, Schrott-Fischer J, Stephen K, Bova S, Chen H, Zheng H, Striessnig J, 2000. Congenital deafness and sinoatrial node dysfunction in mice lacking class D L-type calcium channels. Cell 102: 89–97. Pragnell M, DeWaard M, Mori Y, Tanabe T, Snutch TP, Campbell KP, 1994. Calcium channel bsubunit binds to a conserved motif in the I–II cytoplasmic linker of the a1 subunit. Nature 368: 67–70. Ramkumar V, El-Fakahany EE, 1988. Prolonged morphine treatment increases rat brain dihydropyridine binding sites: possible involvement in development of morphine dependence. Eur J Pharmacol 146: 73–83.

REFERENCES

31

Rampe D, Triggle DJ, 1993. New synthetic ligands for L-type voltage-gated calcium channels. Prog Drug Res 40: 191–238. Raymond CR, Redman CJ, 2006. Spatial segregation of neuronal calcium signals encodes different forms of LTP in rat hippocampus. J Physiol 570: 97–111. Roca G, Aguilar JL, Gomar C, Mazo V, Costa J, Vidal F, 1996. Nimodipine fails to enhance the analgesic effect of slow release morphine in the early phase of cancer treatment. Pain 68: 239–243. Rosse RB, Alim TN, Fay-McCarthy M, Collins JP Jr, Vocci FJ Jr, Lindquist T, Jentgen C, Hess AL, Deutsch S, 1994. Nimodipine pharmacotherapeutic adjuvant therapy for inpatient treatment of cocaine dependence. Clin Neuropharmacol 17: 348–358. Safat P, Boulter J, Hales TG, 2001. Functional properties of Cav1.3 (a1D) L-type channel splice variants expressed in rat brain and neuroendocrine GH3 cells. J Biol Chem 276: 38727– 38737. Sanguinetti MC, Kass RS, 1984. Voltage-dependent block of calcium channel current in calf cardiac Purkinje fibers by dihydropyridine calcium antagonists. Circ Res 55: 336–348. Schulla V, Renstrom E, Feil R, Feil S, Franklin I, Gjinovci A, Jing XJ, Laux D, Lundquist I, Magnuson MA, et al., 2003. Impaired insulin secretion and glucose tolerance in beta cellselective Cav1.2 Ca2þ channel null mice. EMBO J 22: 3844–3854. Scriabine A, 2002. Calcium antagonists and the central nervous system. In: Epstein M (Ed.), Calcium Antagonists in Clinical Medicine. Hanley and Belfus, Inc., Philadelphia, PA. Scriabine A, Schuurman T, Traber J, 1989. Pharmacological basis for the use of nimodipine in central nervous system disorders. FASEB J 3: 1799–1806. Scriabine A, Teasdale GM, Tettenborn D, Young W (Eds), 1991. Nimodipine:. Pharmacological and Clinical Results in Cerebral Ischemia. Springer Verlag, Heidelberg. Sekiya T, Yagihashi A, Asano K, Suzuki S, 2002. Nimodipine ameliorates trauma-induced cochlear neuronal death. Neurol Res 24: 775–780. Shimizu N, Kishioka S, Maeda T, Fukazawa Y, Yamamoto C, Ozaki M, Yamamoto H, 2004. Role of pharmacokinetic effects in the potentiation of morphine analgesia by L-type calcium channel blockers in mice. J Pharmacol Sci 94: 240–245. Sinnegger-Brauns MJ, Hetzenauer A, Huber IG, Renstrom E, Wietzorrek G, Berjukov S, Cavalli M, Walter D, Koscak A, Waldschutz R, et al., 2004. Isoform-specific regulation of mood behavior and pancreatic beta cell and cardiovascular function by L-type Ca2þ channels. J Clin Invest 113: 1430–1439. Smith JW, Watson WP, Stephens DN, Little HJ, 1999. The calcium channel antagonist, nimodipine, decreases operant self-administration of low concentrations of ethanol. Behav Pharmacol 10: 793–802. Soldatov NM, Bouron A, Reuter H, 1986. Different voltage-dependent inhibition by dihydropyridines of human Ca2þ channel splice variants. J Biol Chem 270: 10540–10543. Splawski I, Timothy KW, Sharpe LM, Decher N, Kumar P, Bloise R, Napolitano C, Schwartz PJ, Joseph RM, Condouris K, et al., 2004. Cav1.2 calcium channel dysfunction causes a multisystem disorder including arrhythmia and autism. Cell 119: 19–31. Splawski I, Timothy KW, Decher N, Kumar P, Sachse FB, Beggs AH, Sanguinetti MC, Keating MT, 2005. Severe cardiac arrhythmia caused by cardiac L-type calcium channel mutations. Proc Natl Acad Sci USA 102: 8089–8096.

32

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

Stanley EF, Chan AW, 2005. The calcium channel and the transmitter release site. In: Zamponi G (Ed.), The Voltage-Gated Calcium Channel. Kluwer Academic/Plenum Publishing, New York, pp. 141–153. Striessnig J, Hoda J-C, Koschak A, Zaghetto F, Mullner C, Sinneger-Brauns J, Wild C, Watschinger K, Trockenbacher A, Pelster G, 2004. L-type Ca2þ channels in Ca2þ channelopathies. Biochem Biophys Res Commun 322: 1341–1346. Striessnig J, Koschak A, Sinneger-Brauns MJ, Hetzenauer A, Nguyen NK, Busquet P, Pelster G, Singewald N, 2006. Role of voltage-gated L-type Ca2þ channel isoforms for brain function. Biochem Soc Trans 34: 903–909. Strom TM, Nyakatura G, Apfelstedt-Sylla E, Hellbrand H, Lorenz B, Weber BH, Wutz K, Gutwillinger N, Ruther K, Drescher B, et al., 1998. A L-type calcium channel gene mutated in incomplete X-linked congenital stationary night blindness. Nat Genet 19: 260– 263. Sukriti HD, Pandhi P, 2004. Potentiation of antihyperalgesic activity of diclofenac by nimodipine in a formalin model of facial pain in rats. Methods Find Exp Clin Pharmacol 26: 253–256. Sun JP, Triggle DJ, 1995. Calcium channel antagonists: cardiovascular selectivity of action. J Pharmacol Exp Ther 274: 419–428. Takahashi SX, Miriyala J, Colecraft HM, 2004. Membrane-associated guanylate kinase-like properties of b-subunit required for modulation of voltage-dependent Ca2þ channels. Proc Natl Acad Sci USA 101: 7193–7198. Taylor CP, 2004. The biology and pharmacology of calcium channel a2d proteins. Pfizer Symposium at the 2003 Society of Neuroscience Meeting. CNS Drug Rev 10: 183–188. Theopold HM, 1985. Nimodipine (Bay e 9736): a new concept in the treatment of inner ear diseases. Laryngol Rhinol Otol 64: 609–613. Tobares N, Pedromingo A, Bigorra J, 1989. Nimodipine treatment improves cognitive function in vascular dementia. In: Bergener M, Reisberg B (Eds), Diagnosis and Treatment of Senile Dementia. Springer Verlag, Berlin, pp. 361–365. Triggle DJ, 1989. Structure–function correlations of 1,4-dihydropyridine calcium channel antagonists and activators. In: Hondeghem, LM (Ed.), Molecular and Cellular Mechanisms of Antiarrhythmic Agents. Futura Publishing, Mt. Kiscoe, NY, pp. 269–291. Triggle DJ, 2003. The 1,4-dihydropyridine nucleus: a pharmacophoric template. Mini-Rev Med Chem 3: 137–147. Triggle DJ, 2004. Pharmacology of L-type channels. In: McDonough SI (Ed.), Calcium Channel Pharmacology. Kluwer Academic/Plenum Publishers, New York, pp. 21–62. Triggle DJ, 2006. L-type calcium channels. In: Triggle DJ, Gopalakrishnan M, Rampe D, Zheng W (Eds), Voltage-Gated Ion Channels as Drug Targets. Wiley-VCH, Weinheim, Germany, pp. 100–121. Triggle DJ, 2007. Calcium channel antagonists: clinical uses—past, present and future. Biochem Pharmacol 74: 1–9. Triggle DJ, Langs DA, Janis RA, 1989. Ca2þ channel ligands: structure–function relationships of the 1,4-dihydropyridines. Med Res Rev 9: 123–180. Triggle DJ, Gopalakrishnan M, Rampe D, Zheng W (Eds), 2006. Voltage-Gated Ion Channels as Drug Targets. Wiley-VCH, Mannheim. Trouve R, Nahas GG, 1990. Antidotes to lethal cocaine toxicity in the rat. Arch Int Pharmacodyn Ther 305: 197–207.

REFERENCES

33

Trouve R, Nahas GG, Manger WM, Vinyard C, Goldberg S, 1990. Interactions of nimodipine and cocaine on endogenous catecholamines in the squirrel monkeys. Proc Soc Exp Biol Med 193: 171–175. Tsien RW, Barrett CF, 2005. A brief history of calcium channel discovery. In: Zamponi, G (Ed.), Voltage-Gated Calcium Channels. Kluwer Academic/Plenum Publishers, New York, pp. 27–47. Van Petergem F, Clark KA, Chatelain FC, Minor DL, 2004. Structure of a complex between a voltage-gated calcium channel b-subunit and an a-subunit domain. Nature 429: 671–675. Veng LM, Meshes MH, Browning MD, 2003. Age-related working memory impairment is correlated with increases in the L-type calcium channel protein a1D (Cav1.3) in area CA1 of the hippocampus and both are ameliorated by chronic nimodipine treatment. Brain Res Mol Brain Res 110: 193–202. Wei X-M, Luchowski EM, Rutledge A, Su CM, Triggle DJ, 1986. Pharmacologic and radioligand binding analysis of the actions of 1,4-dihydropridine activator–antagonist pairs in smooth muscle. J Pharmacol Exp Ther 239: 144–153. Welling A, Kwan YW, Bosse E, Flockerzi V, Hofmann F, Kass R, 1993. Subunit-dependent modulation of recombinant L-type calcium channels. Molecular basis for dihydropyridine tissue selectivity. Circ Res 73: 974–980. Welling A, Ludwig A, Zimmer S, Klugbauer N, Flockerzi V, Hofmann F, 1997. Alternatively spliced IS6 segments of the alpha1C gene determine the tissue-specific dihydropyridine sensitivity of cardiac and vascular smooth muscle L-type Ca2þ channels. Circ Res 81: 526– 532. Westenbroek RW, Hoskins L, Catterall WA, 1998. Localization of Ca2þ channel subtypes on rat spinal motor neurons, interneurons and nerve terminals. J Neurosci 18: 6319–6330. Wheeler DG, Barrett CF, Tsien RW, 2006. L-type calcium channel ligands block nicotineinduced signaling to CREB by inhibiting nicotinic receptors. Neuropharmacology 51: 27– 36. Wildemann B, Schulin C, Storch-Hagenlocher B, Hacke W, Dithmar S, Kirchhof K, Jansen O, Breitbart A, 1996. Susac’s syndrome: improvement with combined antiplatelet and calcium antagonist therapy. Stroke 27: 149–151. Xu W, Lipscombe D, 2001. Neuronal Cav1.3a1 L-type channels activate at relatively hyperpolarized membrane potentials and are incompletely inhibited by dihydropyridines. J Neurosci 21: 5944–5951. Yaksh TL, 2006. Calcium channels as therapeutic targets in neuropathic pain. J Pain 7: S13– S30. Yamamoto T, Niwa S, Ohno S, Onishi T, Matsueda H, Koganie H, Uneyama H, Fujita S, Takeda T, Kito M, et al., 2006. Structure–activity relationship study of 1,4-dihydropyridine derivatives blocking N-type calcium channels. Bioorg Med Chem Lett 16: 798–802. Zamponi G (Ed.), 2005. Voltage-Gated Calcium Channels, Kluwer Academic/Plenum Publishers, New York. Zarauza R, Saez-Fernandez AN, Iribarren MJ, Carrascosa F, Adame M, Fidalgo M, Monedero P, 2000. A comparative study with oral nifedipine, intravenous nimodipine and magnesium sulfate in postoperative analgesia. Anesth Analg 91: 938–943. Zhang Z, Xu Y, Song H, Rodriguez J, Tuteja D, Namkung Y, Shin H-S, Chiamvimonvat N, 2002. Functional roles of Cav1.3 (a1D) calcium channel in sinoatrial nodes. Insight gained using gene-targeted null mutant mice. Circ Res 90: 981–987.

34

NEURONAL L-TYPE VOLTAGE-GATED CALCIUM CHANNELS

Zhang Q, Li J-X, Zheng J-W, Liu R-K, Liang J-H, 2003. L-type Ca2þ channel blockers inhibit the development but not the expression of sensitization to morphine in mice. Eur J Pharmacol 467: 145–150. Zhang H, Maximov A, Fu Y, Xu F, Tang T-S, Tkatch T, Surmeier D J, Bezprozvanny I, 2005a. Association of Cav1.3 L-type calcium channels with Shank. J Neurosci 25: 1037–1049. Zhang X, Odom DT, Koo SH, Conkright MD, Canettieri G, Best J, Chen H, Jenner R, Herbolsheimer E, Jacobsen E, et al., 2005b. Genome-wide analysis of cAMP response element binding protein occupancy, phosphorylation and target gene activation in human tissues. Proc Natl Acad Sci USA 102: 4459–4464. Zhang H, Fu Y, Altier C, Platzer J, Surmeier J, Bezprozvanny I, 2006. Cav1.2 and Cav1.3 neuronal calcium channels: differential targeting and signaling to pCREB. Eur J Neurosci 23: 2297–2310. Zhen W, Stoltefuss J, Goldmann S, Triggle DJ, 1992. Pharmacological and radioligand binding studies of 1,4-dihydropyridines in rat cardiac and vascular preparations: stereoselectivity and voltage-dependence of antagonist and activator interactions. Mol Pharmacol 41: 535– 541. Zhou X, Ono H, Ono Y, Frohlic E, 2002. N- and L-type calcium channel antagonist improves glomerular dynamics, reverses nephrosclerosis, and inhibits apoptosis and proliferation in an LNAME/SHR model. J Hypertens 20: 993–1000. Zhulke RD, Bouron A, Soldatov NM, Reuter H, 1998. Ca2þ channel sensitivity towards the blocker isradipine is affected by alternative splicing of the human a1C subunit gene. FEBS Lett 42: 220–224.

2 VOLTAGE-GATED N-TYPE AND T-TYPE CALCIUM CHANNELS AND EXCITABILITY DISORDERS ELIZABETH TRINGHAM1 1 2

AND

TERRANCE P. SNUTCH1,2

Neuromed Pharmaceuticals, Rm 301, 2389 Health Sciences Mall, Vancouver, BC, Canada Michael Smith Laboratories, University of British Columbia, Vancouver, BC, Canada

2.1 INTRODUCTION Calcium (Ca2þ) is a ubiquitous signaling molecule involved in a diverse array of cellular processes ranging from control of membrane excitability to gene transcription, all of which require that intracellular levels be tightly regulated. In particular, voltage-gated Ca2þ channels (Cav) regulate transmembrane fluxes of calcium in response to membrane depolarization. Ca2þ channel currents can generally be grouped into two major classes of Cav channels: high voltage activated (HVA) and low voltage activated (LVA or “T-type”). Based on their pharmacological and biophysical properties, the HVA class can be further subdivided into L-, N-, P/Q-, and R-type channels. The main pore-forming a subunits of the Cav channels are encoded by 10 different genes, CACNA1A, CACNA1B, CACNA1C, CACNA1D, CACNA1E, CACNA1F, CACNA1G, CACNA1H, CACNA1I, and CACNA1S, with a greater variety of subtypes for each class arising from a substantial degree of alternative splicing. Associated with the pore-forming subunit of HVA channels are several accessory subunits, a2d (four genes in mammals), b (four genes in mammals), and g (up to eight genes in mammals), although the precise complement is not well determined biochemically for all of the HVA channel types. Further, HVA channel Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

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FIGURE 2.1 Biophysical properties of high-voltage-activated Cav channels and T-type channels (LVA). (a) Voltage dependence of activation and inactivation of T-type channels is hyperpolarized compared to HVA channels. T-type channels inactivate faster than HVA channels (b), but deactivate (close) more slowly upon repolarization of the membrane potential (c). Adapted from Snutch, 2008.

complexity occurs as a result of the interaction between the main pore-forming HVA Cav subunits and various alternatively spliced forms of the accessory subunits. This complexity in the molecular makeup underlying the Cav channel composition creates a daunting assortment of functional diversity in the physiological roles of Cav channels. As progress has been made in assigning physiological roles to particular Cav channel complexes, it has also unveiled their contributions to numerous pathophysiological conditions beyond the classically defined role of L-type channels in hypertension, angina, and cardiac arrhythmias. More recent interest has focused on understanding the roles of N- and T-type channels, which despite having quite distinct biophysical and pharmacological properties (Fig. 2.1) appear to contribute to the pathophysiology of disorders that are characterized by changes in neuronal excitability. Biophysically, Ntype channels activate at higher potentials than T-type channels, exhibit slower kinetics of activation and inactivation, and recover more slowly from inactivation but deactivate more quickly. Three different genes, CACNA1G, CACNA1H, and CACNA1I, encode the pore-forming subunits of Cav3.1, Cav3.2, and Cav3.3 T-type channels, respectively, whereas N-type (Cav2.2) channels are encoded by a single gene, CACNA1B. Pharmacologically, to date there are no reported subtype-selective T-type channel blockers, which has limited the characterization of these channels in vivo. In contrast, selective peptide blockers with nanomolar affinity to N-type channels are available, providing insight into their roles in neuronal physiology. Despite some pharmacological limitations, the combination of available agents together with molecular genetic animal model and limited clinical data also provide valuable insight into the pathophysiological roles in which both N- and T-type channels are involved.

2.2 PATHOPHYSIOLOGY OF PAIN Major advances in the field of pain have expanded our current understanding of the mechanisms underlying different pain states as well as some of the complexities of the signaling pathways involved in the processing of painful stimuli. One concept that has emerged as a common theme among different pain states, whether arising from neuropathic or inflammatory pain, is the spontaneous and persistent repetitive activation of primary afferent neurons (Kajander and Bennett, 1992; Kajander et al., 1992;

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McLachlan et al., 1993; Devor et al., 1994; Petersen et al., 1996; Ramer et al., 1997; Song et al., 1999; Zhang et al., 1999). These changes in excitability result in exaggerated transmitter release and enhanced postsynaptic excitability of neurons in the dorsal horn, which in turn leads to central sensitization or “windup.” Whether pain arises from damage of tissue as a result of sensitization of sensory terminals by peripheral chemical mediators or nerve damage as in neuropathic pain, both lead to maladaptive changes in central neuronal excitability and hence to behavioral phenotypes such as hyperalgesia and allodynia. Although a number of transmitters are released by primary afferent neurons, at the core of central sensitization is the excitatory neurotransmitter glutamate, the release of which from C and Ad fibers is controlled by Cav channels (Fig. 2.2). During inflammation and neuropathic pain states, both somatic and presynaptic Ca2þ channel activities are elevated leading to an increase in ectopic discharges and

FIGURE 2.2 Primary afferent nociceptive pathway. Following stimulation of primary afferent Ad and C fibers, second-order neurons in the spinal dorsal horn are activated and sensitized by neurotransmitters released into the presynaptic cleft. The second-order neurons ascend as part of the spinothalamic tract where they synapse onto third-order neurons in the thalamus that radiate to the somatosensory cortex where pain is perceived. A subset of neurons transmits proprioceptive information via the dorsal column. N-type calcium channels at primary afferent terminals in the spinal dorsal horn trigger the release of excitatory (glutamate) and modulatory (substance P and CGRP) transmitters onto second-order neurons. In the primary afferent pathway, T-type calcium channels are concentrated at both free nerve endings and DRG cell bodies. Both N-type and T-type calcium channels are also found in second-order spinal neurons as well as in higher brain regions such as the thalamus and cortex. Adapted from Hildebrand and Snutch, 2006.

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glutamate release. It is this increase that results in the activation of glutamate receptors, specifically NMDA receptors through voltage-dependent Mg2þ unblock, which then prolongs neurotransmission and leads to “windup” in spinal nociceptive neurons of the dorsal horn. Further, as Cav channels are also known to fine-tune neuronal Ca2þ dynamics and hence shape action potential waveforms and control firing frequency, they present an interesting opportunity concerning nociceptive therapies. 2.2.1

Role of N-Type Channels in the Pathophysiology of Pain

Since native N-type channels were first identified more than 20 years ago in sensory chick dorsal root ganglion (DRG) using electrophysiology techniques (Nowycky et al., 1985), RNA expression studies have revealed that they are exclusively expressed in neurons of both the central and peripheral nervous systems and in neurally derived cells such as neuroendocrine cells (Dubel et al., 1992). Immunohistochemical studies show that N-type channels are distributed at the subcellular level along dendrites and cell bodies of many neurons and also at a subset of nerve terminals (Westenbroek et al., 1992). Their pivotal presynaptic location suggests that they are likely to be directly involved in synaptic transmission by controlling transmitter release and hence processing of sensory information Koyano et al., 1987. Indeed, the release of neuropeptides from the spinal cord, such as calcitonin gene-related peptide and substance P, is coupled to N-type channels (Holz et al., 1988; Maggi et al., 1990; Santicioli et al., 1992; Evans et al., 1996; Smith et al., 2002). Further, utilizing the peptide N-type Cav blocker w-conotoxin GVIA (w-CTx-GVIA, see below), the direct involvement of N-type channels in regulation of glutamatergic synaptic transmission, a major transmitter in afferent A and C fibers, has been demonstrated between primary afferent neurons and the spinal cord (Gruner and Silva, 1994). Following nerve injury, some inconsistencies have been noted with regard to changes in the expression of N-type channels. For example, N-type immunoreactivity increases in both the small dorsal root ganglion and lamina II of the spinal cord following chronic nerve compression injury (Cizkova et al., 2002), while no change in Cav2.2 mRNA levels is apparent in L5/L6 in a nerve ligation model of neuropathic pain (Luo et al., 2001). In other studies, whole-cell electrophysiological recordings of DRGs following nerve ligation or fura-2 microfluorometry of DRGs from axotomized neurons both demonstrate a reduction in the w-CTx-GVIA-sensitive N-type current (Baccei and Kocsis, 2000; Fuchs et al., 2007). While biochemical evidence implicates N-type channels in the pathophysiology of pain, the precise sensory modalities that N-type channels regulate have been elucidated by peptidic subtype-selective N-type channel blockers. Toxins derived from the venoms of cone snails and hunting spiders have been shown to be potent pain relievers in various animal models of neuropathic and inflammatory pain. One such toxin, w-CTx-GVIA, is derived from the venom of the cone snail, Conus geographus. At nanomolar concentrations, w-CTx-GVIA is a selective and irreversible blocker of N-type channels that acts by occluding the pore of the channel (Olivera et al., 1984, 1987; Wagner et al., 1988; Boland et al., 1994; Feng et al., 2003). Early evidence for the involvement of N-type channels in nociceptive

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responses was demonstrated by autoradiographic studies showing that radiolabeled w-CTx-GVIA binds in a highly localized manner in both the soma and presynaptic terminals of both myelinated and unmyelinated nociceptive primary afferent neurons at the level of superficial laminae I and II of the dorsal spinal cord (Kerr et al., 1988; Gohil et al., 1994). Intrathecal application of w-CTx-GVIA attenuates tactile allodynia and thermal hyperalgesia in the chronic constriction injury, partial sciatic nerve injury, and vincristine models of sensory peripheral neuropathy (Xiao and Bennett, 1995; Yamamoto and Sakashita, 1998; Scott et al., 2002; Fukuizumi et al., 2003a). In the models of inflammatory pain in rats, w-CTx-GVIA reduces hyperalgesia induced by intraplantar injection of carrageenan (Yokoyama et al., 2003) and attenuates the algogenic response of bradykinin and a,b-methylene ATP when injected intraplantarly into mice (Kato et al., 2002). Antinociceptive effects have also been reported with w-CTx-GVIA in the formalin (Murakami et al., 2001, 2004) and capsaicin inflammatory pain models (Sluka, 1997), both of which activate unmyelinated C fibers (Dickenson et al., 1987; Winter et al., 1995; McCall et al., 1996). In a postoperative rat pain model, intrathecal administration of w-CTxGVIA also dose-dependently attenuates incision-induced allodynia (Cheng et al., 2006). In contrast, responses to intrathecal administration of w-CTx-GVIA have provided mixed results in the acute thermal tail-flick antinociception test, with some groups reporting no change in the basal tail-flick latencies (Suh et al., 1997; Scott et al., 2002) while others finding dose-dependent increases in tail-flick latencies in rats and mice (Omote et al., 1996; Lia and Prado, 1999; Fukuizumi et al., 2003a). Spinal nociceptive responses to acute peripheral mechanical stimulation have also implicated N-type channels in increasing threshold responses in the tail and paw pressure tests (Omote et al., 1996; Fukuizumi et al., 2003a). Considering the electrophysiological properties of the neurons involved in sensory signaling, it is interesting to note the commonality among different pain states such as neuropathic and inflammatory pain, namely, that both lead to spontaneous activity in primary afferent fibers (Wall and Devor, 1983; Babbedge et al., 1996; Boucher et al., 2000; Liu et al., 2000; Wu et al., 2001; Djouhri et al., 2006; Rojas-Piloni et al., 2007). This spontaneous activity originating in the peripheral afferents is thought to mediate maladaptive changes in the central nervous system (CNS) and is implicated in sensory abnormalities, including hyperalgesia and allodynia. This peripheral afferent signal is transmitted with a high degree of fidelity to the CNS and studies have shown that Ca2þ influx through N-type channels is critical to both sensitization and plasticity. Extracellular electrophysiological recordings within dorsal horn neurons following spinal nerve ligation show that increased neuronal excitability elicited by application of mechanical, thermal, or electrical stimulation in rats is dose-dependently reduced by application of w-CTx-GVIA to the spinal cord (Matthews and Dickenson, 2001b). In spinal cord neurons that have become excitable following intraarticular injection of either mustard oil or kaolin and carrageenan, w-CTx-GVIA reduces neuronal excitability in both models suggesting a prominent role of N-type channels in afferent C-fiber-mediated hyperexcitability of spinal neurons (Neugebauer et al., 1996; Nebe et al., 1998). Intradermal application of formalin induces a biphasic

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(phase I and phase II) increase in activity in C fibers that corresponds to the biphasic behavioral response. In agreement with its ability to suppress nociceptive behavior, w-CTx-GVIA reduces the spontaneous action potential firing rate in both phases (Dickenson and Sullivan, 1987; McCall et al., 1996; Diaz and Dickenson, 1997). Antinociceptive responses have also been reported for two other N-type channel preferring toxins, w-conotoxin CVID (w-CTx-CVID) and w-conotoxin MVIIA (w-CTx-MVIIA), both of which act by physically occluding the pore of Cav2.2 channels at nanomolar concentrations. w-CTx-MVIIA, derived from Conus magus (Olivera et al., 1985, 1987), reduces the neuropathy-induced allodynia following spinal nerve ligation (Bowersox et al., 1996; Urban et al., 2005) as well as thermal hyperalgesia in both a chronic constriction injury and a partial sciatic nerve injury model (Yamamoto et al., 1998). Mechanical hyperalgesia, assessed following inflammation induced by intraplantar injection of Freund’s complete adjuvant, is also dose-dependently reduced following intrathecal administration of w-CTx-MVIIA (Bowersox et al., 1996; Wang et al., 1998). Interestingly, in the formalin test, both phase I (acute phase) and phase II (tonic phase) are suppressed by w-CTx-MVIIA at doses that are
1000-fold more potent than morphine (Malmberg and Yaksh, 1994, 1995; Bowersox et al., 1996). Similarly, w-CTx-CVID, derived from Conus catus (Lewis et al., 2000; Nielsen et al., 2000), reduces mechanical and tactile allodynia in a spinal nerve ligation model (Scott et al., 2002; Blake et al., 2005) as well as being antinociceptive in the rat Freund’s complete adjuvant model (Smith et al., 2002). Acute nociceptive responses in the rat tail-flick model appear unaffected by either wCTx-CVID or w-CTx-MVIIA (Scott et al., 2002; Blake et al., 2005). Following continuous intrathecal infusion with w-CTx-MVIIA, however, a significant reduction in the response to the hot plate is observed (Malmberg and Yaksh, 1995). Of interest is the observation that although the three subtype-selective w-conotoxins have similar affinity for the N-type channel, they are different in their rank order of potency in attenuating neuropathic pain (Scott et al., 2002). w-CTx-GVIA is the most potent with a three- to fourfold higher potency than w-CTx-MVIIA and w-CTx-CVID, but conversely w-CTx-CVID provides the greatest therapeutic margin, as animals display fewer adverse effects such as serpentine tail movements and body shaking. While this may possibly be explained by actions at off-target sites in the CNS other than N-type channels, as these effects are not noted in the Cav2.2 knockout mice, they support the notion that conotoxins have preferential affinities for different variants of N-type channels. For example, Adams et al. (2003) found that while transmitter release at preganglionic nerve terminals is regulated by N-type channels, w-CTx-CVID but not w-CTx-MVIIA blocks transmitter release. To address chronic pain conditions in patients refractory to current therapies, including morphine, the administration of a synthetic version of w-CTx-MVIIA, currently marketed as Prialt, has recently become available. Prialt was approved by the Food and Drug Administration (FDA) in 2005 to treat severe chronic cancer or AIDS pain and in some cases has resulted in complete or near-complete pain relief in patients whose symptoms were previously unmanageable even by intrathecal morphine (Brose et al., 1997; Mathur, 2000; Staats et al., 2004; Wallace, 2006). These results indicate that the animal models of pain used to evaluate the role of N-type

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channels in somatosensory processing of pain were highly predictive of human clinical outcome. The lack of tolerance of Prialt in humans was also predicted in animal models of pain as continuous intrathecal infusion of w-CTx-MVIIA in rats minimally developed tolerance after 7 days (Malmberg and Yaksh, 1995). Side effects with Prialt clinical use have been reported and can be present in the form of hallucinations, cognitive impairment, and alterations in mood and consciousness, thereby providing the opportunity for improved N-type channel blockers that more specifically target those channels resident in sensory pathways (Penn and Paice, 2000; Snutch, 2008). In support of in vivo behavioral tests using pharmacological tools to assign sensory modalities regulated by N-type channels, mice strains in which the Cav2.2 N-type has been genetically removed have for the most part substantiated the results using w-conotoxins. The data from three knockout strains from different laboratories clearly show the involvement of the N-type channels in the development of allodynia and hyperalgesia associated with neuropathic and inflammatory pain (Kim et al., 2001; Saegusa et al., 2001, 2002). However, for acute pain, the role of N-type channels is somewhat ambiguous as results are inconsistent among the labs. Interestingly, the three independent Cav2.2-deficient mouse strains exhibit surprisingly few deleterious effects despite the wide CNS distribution and known role of N-type channels in synaptic transmission. The most marked behavioral alteration related to CNS function is decreased anxiety, whereas in the autonomic nervous system, sympathetic nerve dysfunction is observed as a reduction in the baroreflex and elevated heart rate and blood pressure (Ino et al., 2001). As the assortment of splice variants encoding Cav channels is being unraveled, the precise alternatively spliced isoform of Cav2.2 expressed in sensory neurons has recently unfolded (see Lipscombe et al., Chapter 3 in this volume). A Cav2.2 splice isoform containing exon 37a is preferentially expressed in capsaicin receptorexpressing nociceptive neurons whereas the exon 37b variant is ubiquitously expressed (Bell et al., 2004). Of particular interest are the recent findings by Altier and colleagues showing that when splice isoform-specific small interfering RNA (siRNA) against exon 37a Cav2.2 channels is injected intrathecally, this results in a reduction in both thermal and mechanical hyperalgesia in inflammatory and neuropathic pain models, as well as in basal thermal nociception. Tactile neuropathic allodynia is, however, equally mediated by exon 37a- and 37b-containing neurons (Altier et al., 2007). In addition to these in vivo findings is the identification of a distinct mode of G-protein-mediated voltage-independent inhibition unique to Cav2.2e[37a] (Raingo et al., 2007). This is in contrast to the well-defined voltage-dependent G-protein inhibition of Cav2.2 channels promoted by the liberation of Gbg from Gi/o heteromers following agonist–receptor binding (i.e., morphine) (Herlitze et al., 1996; Ikeda, 1996). Although both exon 37a- and 37b-containing Cav2.2 splice isoforms undergo varying degrees of voltage-dependent inhibition, voltage-independent inhibition is only present in the Cav2.2 channels containing the 37a variant. The signaling pathway coupling G-protein-coupled receptors (GPCRs) to voltage-independent inhibition of Cav2.2e[37a] is mediated by the Ga subunit with downstream activation of a tyrosine kinase, which phosphorylates a single tyrosine residue (Y1747) in exon 37a. The

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physiological significance that has been speculated of this bifurcating pathway is that it maximizes the efficiency of inhibition by using both Ga and Gbg subunits from Gi/o heteromers, while simultaneously providing adaptive changes to Ca2þ influx depending on the level of neuronal activity (Ikeda and Dunlap, 2007). This will no doubt stimulate interest in generating a Cav2.2e[37a]-selective blocker that might have an increased therapeutic index while narrowing the risk of unwanted side effects. 2.2.2

Role of T-Type Channels in the Pathophysiology of Pain

Despite being identified in rat and chick dorsal root ganglion over 20 years ago (Carbone and Lux, 1984a, 1984b; Nowycky et al., 1985), a prominent role of T-type channels in the pathophysiology of pain has only recently emerged. This has been attributed in part to the discovery of three distinct subunit genes, CACNA1G (Cav3.1), CACNA1H (Cav3.2), and CACNA1I (Cav3.3), that encode T-type channels, as well as the existence of several classes of nonselective T-type channel blockers that have been used to tease out physiological and pathophysiological roles. In comparison to N-type channels, T-type calcium channels are much more widely expressed. T-type calcium channel currents are found in many tissues and cell types, including both central and peripheral neurons, heart, adrenal glands, kidney, smooth muscle, embryonic skeletal muscle, pituitary, pancreas, retina, and testes (PerezReyes, 2003). In the somatosensory pathway, the expression of T-type channels is localized to the dorsal root ganglion, dorsal horn spinal neurons, and the thalamus. In the dorsal root ganglion neurons, characterization of these primary afferents using electrophysiological techniques determined that T-type channels are exclusively expressed in small- and medium-sized DRG neurons but are essentially devoid of expression in the large DRG neurons (Schroeder et al., 1990; Scroggs and Fox, 1992; Shin et al., 2003). More recently, a novel subset of small capsaicin-sensitive DRG neurons called “T-rich” neurons has been identified that express only T-type channels with little contribution of the calcium conductance attributed to HVA channels (Nelson et al., 2005). Generally, the nociceptive C and Ad fibers are considerably smaller than Aa and Ab fibers that conduct primarily proprioceptive and tactile information (Harper and Lawson, 1985a, 1985b), which supports the contribution of T-type channels in the pathophysiology of pain. Of the three genes known to encode T-type channels, in situ hybridization and reverse-transcription PCR studies have identified the Cav3.2 subtype as the most prominent subtype in DRG neurons, whereas Cav3.3 expression is lower and Cav3.1 is virtually undetected in small to medium DRGs (Talley et al., 1999; Bourinet et al., 2005). In another subset of medium-sized DRG neurons, the Cav3.2 T-type channel is found to be present in D-hair cell mechanoreceptors (Shin et al., 2003). At the level of the spinal cord, in situ hybridization studies have demonstrated that the dorsal horn neurons in the superficial lamina also express Cav3.2 (Talley et al., 1999). In contrast to the predominant role of the Cav3.2 subtype of T-type channel in the primary and secondary order neurons of the somatosensory pathway, the thalamus expresses all three subtypes, albeit differentially expressed in subsets of thalamic neurons. Cav3.1 is predominantly expressed in

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the thalamocortical (TC) neurons, which relay information from the periphery to the cortex, and Cav3.2 and Cav3.3 are found in the reticular thalamic neurons that provide inhibitory input to TC neurons (Talley et al., 1999). In the absence of subtype-selective blockers for Cav3.1-, Cav3.2-, or Cav3.3containing T-type channels, knockdown and knockout studies have provided the greatest insight into the sensory modalities conveyed by the individual isoforms. Intrathecal injection of antisense oligonucleotides against each subtype identified Cav3.2 T-type channel as having a major pronociceptive role, whereas antisense directed at Cav3.1 and Cav3.3 did not significantly affect the nociceptive behavioral responses in rats (Bourinet et al., 2005). Cav3.2 T-type channels have effects on both acute and chronic pain as knockdown of Cav3.2 results in antiallodynic and antihyperalgesic effects in rats with a chronic constriction nerve injury and is antinociceptive in na€ıve rats when a noxious pressure is applied to the paw or a noxious thermal stimulus is applied to the tail. Concomitantly, whole-cell recordings reveal that T-type currents are reduced by 70–90% in small- to medium-sized DRGs, following a 50% reduction in Cav3.2 mRNA levels. In contrast, in mice lacking the Cav3.2 subtype of T-type Ca2þ channel, thermal hyperalgesia and mechanical allodynia are not reduced following spinal nerve ligation (Choi et al., 2007). Responses to acute pain, such as noxious mechanical (tail clip) and thermal (tail flick and hot plate) stimuli in na€ıve mice, are, however, reduced in Cav3.2deficient mice as are responses to intradermal capsaicin and formalin and visceral injections of acetic acid and MgSO4, both of which induce abdominal writhing. Although the role for Cav3.1 channels in processing of sensory inputs is predicted to be minimal as expression levels in the DRGs are extremely low or undetected, Cav3.1 knockout mice are unexpectedly found to have increased visceral pain response when induced by intraperitoneal injections of acetic acid or MgSO4 (Kim et al., 2003). In addition to increased visceral pain responses in the mutant mice, there are also changes in the firing patterns in the ventroposterolateral (VPL) thalamocortical neurons. Administration of acetic acid in wild-type mice increases both single spikes and bursts of spike activity in VPL neurons, but in the Cav3.1 knockout mice, only the frequency of the single spikes increases. However, the burst activity is not observed either prior to or following intraperitoneal administration of acetic acid in Cav3.1 knockout mice. This change in pattern of firing is likely to decouple the TC neurons from efficiently activating neurons in the nRT, which in turn provide the inhibitory feedback to the VPL neurons to induce burst firing. By preventing burst firing, the “sensory gate” is removed and sensory signals are able to reach the somatosensory cortex. This antinociceptive role of the thalamus is not thought to be effective at controlling acute pain responses, which are mediated by local reflexes in the periphery. Supporting this notion that Cav3.1 does not contribute to acute pain pathways is a lack of any difference in pain responses to thermal (tail flick and paw withdrawal) or mechanical stimuli (von Frey) in both Cav3.1 mutant and wild-type littermates. However, in the complete Freund’s adjuvant model of chronic inflammation, a model that involves central sensitization, hyperalgesia scores also do not differ in wild-type and mutant mice following intraplantar injection.

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Although there are no high-affinity subtype-selective T-type channel blockers available to support in vivo knockout and knockdown studies, there are nonselective compounds known to inhibit these channels, which have provided insight into the role of T-type channels in processing nociceptive signals. Ethosuximide, a known antiepileptic drug, inhibits cloned Cav3.1, Cav3.2, and Cav3.3 (
0.3–1 mM) at therapeutic plasma levels (
0.3–0.7 mM; Gomora et al., 2001) and also dosedependently suppresses both thermal hyperalgesia and mechanical allodynia following L5/L6 spinal nerve ligation when administered i.p. (Dogrul et al., 2003). This is in contrast with the lack of effect seen when ethosuximide is injected intrathecally, suggesting a peripheral mechanism of action. Similarly, no effect is seen when injected intraplantarly into the injured paw, suggesting that ethosuximide likely acts in the periphery at the level of the DRGs and not at sensory nerve endings. In other neuropathy models, paclitaxel- and vincristine-induced models of neuropathy, ethosuximide injected i.p. produces significant reversal of mechanical allodynia and hyperalgesia (Flatters and Bennett, 2004). Barton et al. 2005 also report antinociceptive effects of ethosuximide in both the early and late stages of the formalin test and in the acute tail-flick test as well as reversal of capsaicin-induced mechanical allodynia following i.p. injection. In contrast, intrathecal administration of ethosuximide fails to affect the nociceptive responses in the rat formalin test (Cheng et al., 2007). In addition to the behavioral studies, during in vivo recordings from single neurons in the dorsal horn in which ethosuximide is directly applied to the spinal cord, the responses of the dorsal horn neurons to mechanical and thermal stimuli were suppressed in sham-operated and spinal nerve-ligated animals (Matthews and Dickenson, 2001a). Two other antiepileptic drugs that are structurally diverse from ethosuximide and for which T-type inhibition has been reported are zonisamide and phenytoin. In cultured neurons, 50 mM zonisamide inhibits
40% of the T-type current in neuroblastoma cells (Kito et al., 1996) and 60% at 500 mM in cerebral cortex neurons (Suzuki et al., 1992). Interestingly, this compound suppresses neuropathy-induced thermal hyperalgesia, but not mechanical allodynia, in the Bennett chronic constriction rat model (Bennett and Xie, 1988; Hord et al., 2003) and is reported to provide relief in humans with refractory neuropathic pain (Guay, 2003; Takahashi et al., 2004) as well as prophylaxis for migraine patients (Drake et al., 2004). Phenytoin, which is used clinically in humans to treat neuropathic pain (McCleane, 1999; Finnerup et al., 2005), blocks Cav3.1 and Cav3.2 (140 and 8.3 mM, respectively; Todorovic et al., 2000) expressed in human embryonic kidney (HEK) cells, as well as native T-type channels in DRG neurons (IC50 ¼
8.3 mM) and N1E-115 neuroblastoma (3–100 mM; Matsuki et al., 1984; Twombly et al., 1988). Though not studied in wide variety of animal models, phenytoin dose-dependently reduces bradykinin-induced pain when applied subcutaneously (Foong and Satoh, 1983); however, in acute models, i.p. administration of phenytoin is superior at reducing thermal pain in pawwithdrawal test than mechanical pain as measured in the tail-pressure test (Sakaue et al., 2004). Some of the most potent T-type channel blockers are comprised of neuroleptics used to treat a variety of psychiatric disorders, including schizophrenia, Tourette’s

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disorder, and obsessive-compulsive disorder. In particular, pimozide inhibits in a nonpreferential manner Cav3.1, Cav3.2, and Cav3.3 with IC50 values of
50 nM (Santi et al., 2002). Although not extensively tested in a variety of pain models, pimozide when administered at low doses in the mouse formalin test is not highly efficacious (Saddi and Abbott, 2000), yet in humans it suppresses the pain associated with trigeminal neuralgia (Lechin et al., 1989; Green and Selman, 1991). Another compound well known for its inhibition of the T-type channels as well as HVA channels (P/Q-, R-, and L-types) is mibefradil, which inhibits cloned and native T-type channels with IC50 values ranging from
0.1 to 4.7 mM depending on the assay conditions and cell type (Jim
enez et al., 2000; Martin et al., 2000). In contrast to ethosuximide, when mibefradil is administered both intraperitoneally and injected directly into the injured limb of rats, it effectively reduces the tactile allodynia induced by spinal nerve ligation of L5/L6 (Dogrul et al., 2003). A reduction in thermal hyperalgesia is also reported following i.p. administration. Of relevance is the fact that mibefradil does not cross the blood–brain barrier when administered systemically and also does not suppress neuropathic behaviors upon direct i.t. administration, again suggesting a peripheral mechanism of action (Ertel et al., 1997). In the capsaicin assay, the results were mixed in that systemic (i.p) administration has no effect on allodynia, yet when injected intracisternally (i.c.), capsaicin-induced mechanical allodynia is dose-dependently reduced (Barton et al., 2005). In a postoperative pain model involving an incision into the paw, no antiallodynic action is observed following i.t. administration of mibefradil (Cheng et al., 2006). However, mibefradil inhibits both phases of the formalin response when injected i.p. or i.t., but is ineffective in suppressing acute nociceptive responses in the tail-flick reflex when administered i.p. (Barton et al., 2005; Cheng et al., 2007). Also noted is a lack of effect on an acute nociceptive stimulus (tail-flick test) when mibefradil is administered following i.t. administration (Dogrul et al., 2001; Barton et al., 2005). Another class of compounds that inhibit native T-type channels in DRG neurons is 5a-reduced neuroactive steroids, a class of compounds also known to potentiate GABAA ligand-gated channels. Of the various 5a-reduced neuroactive steroids examined, (3b,5a,17b)-17-hydroxyestrane-3-carbonitrile (ECN) was identified as inhibiting T-type channels in DRGs (IC50 300 nM) but being devoid of effects on GABAA receptors (Todorovic et al., 1998). To assess the peripheral role of T-type channels in nociceptive processing of acute noxious thermal stimulus to the paw, ECN injected intradermally dose-dependently produces analgesia as seen by the increase in latency to paw withdrawal (Pathirathna et al., 2005a). In rats with neuropathic pain (Bennett and Xie, 1988), intradermal injection of ECN alleviates both thermal and mechanical hyperalgesia in ligated rats and thermal and mechanical nociception in sham-operated rats (Pathirathna et al., 2005b). This antinociceptive effect on the thermal stimulus in ligated and sham-operated rats is not reversed by bicuculline, a GABAA antagonist, suggesting that the effect is most likely due to T-type channel inhibition. A novel role of peripheral T-type channels in amplifying nociceptive signals was unveiled when the effects of redox agents were examined on cloned and native T-type

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channels. Todorovic et al. (2001) found that the endogenous reducing agent, L-cysteine, augments T-type currents by 130% in small DRG neurons and in recombinant Cav3.2 channels by a similar amount and is reversed by the oxidizing agent 5,50 -dithio-bis-(2-nitrobenzoic acid) (DTNB). The effects are mimicked by the reducing agent dithiothreitol (DTT). Conversely, the oxidizing agent DTNB, when applied alone, inhibits the peak T-type calcium channel current in small DRG neurons and Cav3.2 channels expressed in HEK cells, both by about
50%. When applied intradermally into the ventral side of the paw, the reducing agents decrease pawwithdrawal latencies (PWLs), while the oxidizing agent prolongs PWLs. Cysteine also produces a hyperalgesic response to noxious mechanical stimulus, while DTNB is analgesic and reverses the effect of L-cysteine. In providing further evidence that T-type channels boost nociceptive signals, mibefradil reverses the thermal hyperalgesic response of L-cysteine and DTT and enhances the analgesic effect of DTNB. Similarly, reducing cysteine analogues, L-cysteine, D-cysteine, and D,L-homocysteine, induce potent dose- and time-dependent hyperalgesia and conversely endogenous oxidizing cysteine analogues, L-cysteine, D-cysteine, and D,L-homocysteine, induce potent dose- and time-dependent analgesia in an acute model of thermal peripheral nociception in intact rats (Pathirathna et al., 2006). In a neuropathic pain model induced by chronic constriction of the sciatic nerve, both L-cysteine and DTT increase the thermal hyperalgesic response in both nerve-injured and sham-operated rats, while DTNB reduces the neuropathy-induced thermal hyperalgesia and produces analgesia in sham-operated rats (Todorovic et al., 2004). Both DTNB and mibefradil are able to abolish the L-cysteine-induced increase in thermal hyperalgesia rats with neuropathic pain and in sham-operated rats. At the cellular the level, the mechanisms underlying changes in neuronal excitability that lead to the development of allodynia and hyperalgesia are not well understood (Kajander and Bennett, 1992; Kajander et al., 1992; McLachlan et al., 1993; Devor et al., 1994; Petersen et al., 1996; Ramer et al., 1997; Song et al., 1999; Zhang et al., 1999.). Recent molecular and behavioral pharmacology data have provided a strong linkage for the contribution of T-type channels to the pathological perceptions of pain, but the biophysical properties of T-type channels themselves provide even further evidence that they contribute to hyperexcitability observed in DRGs after peripheral trauma or inflammation. Under normal physiological conditions, T-type channels are capable of regulating the pacemaker activity of neurons, contributing to rebound burst firing and oscillatory behavior and hence making them likely contributors to the hyperexcitability of primary afferent neurons in neuropathic and inflammatory pain states. One of the first studies investigating the involvement of T-type channels in processing sensory information was by White et al. (1989) who reported that Ca2þ influx by LVA channels produces an afterhyperpolarizing potential in dorsal root ganglion that triggers a burst of action potentials, which is abolished by 100 mM Ni2þ. Supporting evidence using the “Trich” subset of DRGs that express a high density of T-type channels showed that the endogenous redox agent, L-cysteine, which promotes an increase in the current amplitude and shifts the gating properties, also lowers the nociceptor excitability threshold and induces burst firing (Nelson et al., 2005). Also in the D-hair

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mechanoreceptor neurons, a role of Cav3.2 T-type channels in inducing a slow depolarization has been identified, which leads to a lowering of the voltage threshold for the generation of action potentials and hence an increase in neuronal excitability (Dubreuil et al., 2004). In a streptozotocin model of diabetic neuropathy, the T-type currents were shown to have increased by twofold in medium-sized DRGs with a concomitant depolarizing shift in the steady-state inactivation, which results in a greater “window current,” suggesting that T-type channels can activate more readily at physiological membrane potentials (Jagodic et al., 2007). Interestingly, Jagodic et al. also identified a newly expressed T-type channel that only partially inactivates at a membrane potential of 40 mV, a voltage that completely inactivates the T-type channels in DRGs from control rats, and likely contributes to the reduced threshold for burst firing in DRGs neurons from diabetic rats. In contrast, T-type channel currents in DRGs were reduced after neuropathic injury was induced by chronic constriction of the sciatic nerve or axotomy of L5 neurons in rats (McCallum et al., 2003; Hogan, 2007). Also observed in these DRGs following the induction of neuropathic pain is an increase in the rate of deactivation, which also supports the loss of T-type channels that deactivate slowly in a voltagedependent manner. As the ability of neurons to fire repetitively requires that there is sufficient influx of Ca2þ to activate Ca2þ-activated Kþ channels that in turn hyperpolarize the cell, a reduction in Ca2þ at the entry during the slow deactivation mode at the end of an action potential may contribute to increased excitability of the DRGs from injured rats. Ultimately it will be interesting to determine whether the ability of T-type channel blockers to provide pain relief is dependent solely on their affinity of inhibiting Cav3.2 versus Cav3.1 and Cav3.3 channels or some combination thereof. Although the available evidence is suggestive of a peripheral mechanism of action, whether the efficacy is enhanced by the CNS-penetrant compounds also needs to be determined. Given the critical ability of Cav3.1 to alter the sensory gate in the thalamus, the effects of modulators on these central channels, which remain to be fully elucidated, may be quite important.

2.3 CONTRIBUTIONS OF T-TYPE CHANNELS TO THE PATHOPHYSIOLOGY OF EPILEPSY Epilepsy is a complex disorder of spontaneous recurrent seizures characterized by neuronal hyperexcitability leading to hypersynchronization of neural networks. Phenotypically, seizures present themselves in different forms depending on the site of origin and subsequent recruitment of additional CNS structures. Although the cellular substrate(s) underlying the genesis of seizure activity are largely unknown, genetic, pharmacological, and physiological evidence all implicate an involvement of T-type calcium channels. The involvement of T-type channels in seizure disorders is primarily implicated in idiopathic generalized epilepsies (IGEs) with evidence arising from studies using transgenic and mutant animals as well as population analysis studying linkages to genetic mutations in humans.

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Of the many IGEs, T-type channels have been shown to be critical for seizure activity in absence epilepsy. Absence epilepsy is characterized by a brief and sudden loss of consciousness, which is temporally correlated with 3 Hz bilateral synchronous spike-wave discharges (SWDs) involving the thalamocortical circuitry (Williams, 1950; Niedermeyer and Primary, 1996). Specifically, rebound burst firing in neocortical cells, thalamic reticular neurons, and thalamocortical relay neurons is known to be evoked by low-threshold Ca2þ potentials, which are thought to give rise to SWDs (Llin
as and Jahnsen, 1982; Deschênes et al., 1984; Jahnsen and Llinas, 1984; Coulter et al., 1989a; Huguenard and Prince, 1992; De la Pena and Geijo-Barrientos, 1996; Destexhe and Sejnowski, 2002). In rats, the mRNAs of all three T-type isoforms are expressed in the thalamocortical pathway, though differentially in thalamus and neocortex (Talley et al., 1999). Cav3.2 and Cav3.3 are expressed in the thalamic reticular nucleus, whereas Cav3.1 is dominantly expressed in the thalamocortical neurons. More recently, splice variants of Cav3.1 have been differentially localized in the thalamic circuitry, which may account for the divergent burst firing in TC and GABAergic interneurons (Broicher et al., 2007a). In the neocortex, Cav3.1 and Cav3.3 mRNA levels are diffusely distributed through most layers of the cortex. Conversely, Cav3.2 mRNA is regionally restricted to layer 5 cortical pyramidal neurons, with little expression in other cortical areas. Of interest is that although the neural networks involved in SWDs all express one or more T-type channel isoform, the initiation and generalization has only recently been demonstrated as arising from the cortex. Meeren et al. (2002) have demonstrated through nonlinear association analysis that SWDs originate from the perioral region of the somatosensory cortex of Wistar Albino Glaxo rats from Rijswijk (WAG/Rij), a genetic model of absence epilepsy. Multisite field potential recordings from other cortical areas consistently lag behind the perioral region, with the cortical focus leading the thalamus during the first 500 ms. More in-depth electrophysiological studies using in vivo intracellular recordings from genetic absence epilepsy rats from Strasbourg (GAERS) demonstrate that the neural substrate involved in the initiation and intracortical propagation of ictal activity leading to generalization of related thalamic nuclei is found in layer 5/6 of the perioral somatosensory cortex (Polack et al., 2007). These neurons display pronounced hyperactivity relative to more superficial cortical neurons in that their membrane potentials are more depolarized and show distinctive interictal and preictal 9–11 Hz oscillations as well as enhanced bursting activity. It is postulated that these changes in layer 5/6 neurons create local oscillations (Silva et al., 1991) that lead to the generation of SWDs that propagate secondarily and intra- and interhemispherically throughout the somatosensory cortex and other cortical regions and the thalamic nuclei. Synchronization of SWDs in the thalamocortical and corticothalamic neurons sets into motion a unified oscillatory network that is driven by the ictogenic properties of the cortical neurons while the thalamocortical neurons provide resonant circuitry to sustain the activity. In support of the “cortical focus theory” of absence epilepsy, infusion of ethosuximide (a first choice antiabsence drug) into the perioral region of GAERS animals but not into the thalamus immediately reduces SWDs (Sherwin,

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1989; Richards et al., 2003; Manning et al., 2004). Actions of ethosuximide on native T-type currents from isolated thalamocortical neurons and neurons from the thalamic reticular nucleus have, however, been mixed, with some groups demonstrating up to a 40% reduction in current amplitude at therapeutic concentrations (Coulter et al., 1989; Huguenard and Prince, 1994; Huguenard, 2002) while other studies observing no inhibition (Pfrieger et al., 1992; Leresche et al., 1998). More recently, a study on cloned channels confirms that ethosuximide inhibits cloned Cav3.1, Cav3.2, and Cav3.3 channels (IC50s
0.3–1 mM) at therapeutic plasma levels (
0.3–0.7 mM; Gomora et al., 2001). Further evidence providing insight into the role of T-type channels in the generation of SWDs and absence-like seizures is from Cav3.1 knockout mice (Kim et al., 2001). Not only are Cav3.1 KO mice resistant to baclofen-induced 3–5 Hz spike-wave discharges as measured by EEG, intracellular recordings from thalamocortical neurons show an absence of rebound burst firing action potentials in thalamic slices when injected with negative current. In freely moving mice, field potential recordings from the ventroposteromedial and ventroposterolateral nuclei also show that responses to baclofen-mediated intrathalamic oscillations, which are required for SWD discharges, are also diminished. On the other hand, low doses of bicuculline, which evokes SWDs originating from the cortex (Steriade and Contreras, 1998), when injected into Cav3.1 KO mice are capable of generating SWDs in both the thalamus and cortex. Similarly, Cav3.1 KO mice are not resistant to 4-aminopyridine (4-AP)induced tonic–clonic seizures. While these experiments highlight a critical role of Cav3.1 in generating absence-like seizures, the lack of protection in other models may be indicative of the dominant involvement of either Cav3.2 or Cav3.3 in the cortex for bicuculline-induced SWDs and mechanistically different pathways for 4-AP-induced seizures. Cross-breeding Cav3.1 KO mice with other models of absence seizures, such as lethargic (b4lh/lh), tottering (a1Atg/tg), or stargazer (g2stg/stg) mutant mice models or mice harboring a null mutation for the pore-forming Cav2.1 subunit of P/Q-type channels, results in strongly or completely suppressed cortical SWD paroxysmal activities (Song et al., 2004). Interestingly, in all these murine models, T-type channel currents are elevated in thalamocortical neurons by up to
50%, although no alteration in mRNA levels are apparent (Zhang et al., 2002; Nahm et al., 2005). In GAERS animals, elevated increases in reticular thalamic T-type channel currents (
55%) have also been reported with a concomitant increase in Cav3.1 and Cav3.2 mRNA levels in the ventral posterior thalamic relay nuclei and thalamic reticular nucleus, respectively (Tsakiridou et al., 1995; Talley et al., 2000). More recently, a homozygous, missense, single nucleotide (G–C) mutation has been identified in the Cav3.2 gene from GAERS, though the functional effects are unknown (Kyi et al., 2006). However, in Cav2.1// Cav3.1/þ mice, T-type channel currents are decreased by
25%, yet animals are still capable of generating SWDs indicating that baseline levels, but not necessarily augmented levels, of T-type channel currents are sufficient for the genesis of spontaneous SWD activity. As protection against absence seizures was afforded by deletion of Cav3.1, it will be interesting to determine whether complete and selective pharmacological inhibition of Cav3.1 channels provides complete

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VOLTAGE-GATED N-TYPE AND T-TYPE CALCIUM CHANNELS

suppression of SWDs in these genetic models and pharmacological models of absence seizures. In humans, the role of T-type channels as a susceptibility gene leading to the pathophysiology of absence epilepsy has been speculated due to the known function of T-type channels in the thalamic and cortical physiology. All three subtypes, Cav3.1, Cav3.2, and Cav3.3, have been examined in patients with childhood absence epilepsy (CAE) and other forms of IGEs. Population analyses studying linkages to genetic mutations in humans have identified numerous missense mutations in the Cav3.2 gene. Chen et al. (2003a) reported 12 mutations in 14 patients of Chinese Han ethnicity that were not observed in 230 control patients. In some patients, two or more mutations were identified and numerous polymorphisms were reported in both patients and seizure-free individuals. In an expanded study, which included patients with CAE, juvenile myoclonic epilepsy, and febrile convulsions, three new missense mutations have been identified, though they were also found in the control group (Heron et al., 2004). More recently, 28 new variants have been identified in the Cav3.2 gene from the Chinese Han population of which only some were found in CAE patients (Liang et al., 2006). In a study attempting to investigate whether common polymorphisms in the Cav3.2 gene are associated with CAE in the Chinese Han population, carriers with three different polymorphisms were identified as at a higher risk of developing CAE than noncarriers (Liang et al., 2007). In a predominantly Caucasian population, over 100 variants of the Cav3.2 gene have been found in patients with IGEs as well as temporal lobe epilepsy (Heron et al., 2007). In contrast, in an evaluation of Caucasian European patients with CAE, linkage analysis was unable to detect any of the Chinese variants in 220 patients (Chioza et al., 2006). The human Cav3.2 mutations are located mainly in exons 6–12 encoding the domain I–II linker region. Exogenous expression of the mutations introduced into rat and human Cav3.2 channels reveals that in some instances channel biophysical properties are altered while other changes are “biophysically silent” (Khosravani et al., 2004, 2005; Vitko et al., 2005; Peloquin et al., 2006). Alterations in the biophysical properties mainly result in gain-of-function phenotypes as seen by a more depolarized steady-state inactivation and hyperpolarized voltage dependence of activation. Further insight into the possible roles of the “biophysically silent” mutations and SNPs in loop I–II was revealed when deletion of this region was found to increase Cav3.2 plasma membrane expression with a predicted overactivity to occur in neurons (Vitko et al., 2007). Therefore, the effect of the gain of function as a result of changes in biophysical properties on the Cav3.2 channels as well as biophysically silent mutations that may serve to promote increased plasma membrane expression both result in increased T-type channel activity. A study investigating whether Cav3.1 channels are involved in the etiology of IGEs has proposed a linkage to the CACNA1G gene (Singh et al., 2007). However, while 13 variants were identified in 123 Japanese and Hispanic patients with IGEs, many were found in both patients and control individuals. In the Han Chinese population, no Cav3.1 mutations were identified in patients with CAE, but six single nucleotide polymorphisms (SNPs) were found (Chen et al., 2003b). Overall, the distribution of SNPs was not significantly different in control and CAE patients in the Chinese Han

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population and therefore the SNPs are not considered important for susceptibility to nonconvulsive seizures (Wang et al., 2006). Though the pathophysiological role of T-type channels has primarily focused on IGEs, they are also likely to be involved in other seizure types as RNA expression studies have shown that Cav3.1 and Cav3.3 are prominently expressed in the human CNS, including structures known to generate seizure activity, such as the thalamus, cortex, hippocampus, and amygdala. In situ hybridization studies in rat brain have also revealed diffuse mRNA expression of Cav3.1, Cav3.2, and Cav3.3 in all brain structures (Craig et al., 1999; Talley et al., 1999). Interestingly, at the subcellular level subtypespecific polyclonal antibodies against Cav3 channel proteins indicate that the three T-channel subtypes are differentially localized in the soma and dendrites (McKay et al., 2006). Cav3.1 immunolabeling is prominent in the soma and proximal region of the dendrites, while somatic and proximal middendritic regions largely express Cav3.2. Distribution of Cav3.3 is further distinct in that expression is found in the soma as well as extended throughout the arborization of the dendrites in selective neurons. In concordance with this immunohistochemical study are previous electrophysiological studies that have identified T-type currents in both the somatic and dendritic compartments of central neurons (Karst et al., 1993; Markram and Sakmann, 1994; Magee and Johnston, 1995; Mouginot et al., 1997; Jung et al., 2001; Isope and Murphy, 2005). Two particular animal models of seizures have substantiated the potential role of T-type channels in seizure types other than IGEs. In the pilocarpine-induced status epilepticus rat model, a model of human complex partial seizures with secondary generalization, a threefold increase in T-type tail current density is observed in hippocampal neurons with a concomitant 54% increase in the number of neurons with intrinsic burst firing, which normally fire in a regular mode; this was inhibited by Ni2þ with an IC50 of 27 mM (Su et al., 2002). In a focal model of epilepsy induced by kindling of the rat hippocampus, patch-clamp recordings from in situ slices show an
80% increase in T-type channel currents in hippocampal neurons compared to recordings from control rats (Faas et al., 1996). These elevated T-type channel currents are still present 6 weeks after the last kindling stimulation and support a role of T-type channels in epileptogenesis. Definitive pharmacological proof-of-concept studies confirming that T-type calcium channels represent important drug targets for the treatment of different epilepsies have been hampered by the lack of high-affinity subtype-selective blockers. Besides ethosuximide, which is used to treat patients with absence epilepsy and inhibits both native (0.2–24 mM) and cloned T-type channels (Cav3.1, Cav3.2, and Cav3.3;
0.3– 1 mM) at therapeutically relevant concentrations of
0.3–0.7 mM (Coulter et al., 1989b; Huguenard and Prince, 1994; Gomora et al., 2001; Huguenard, 2002), only a handful of other AEDs with activity at T-type Ca2þ channels have been identified. Valproate, a broad spectrum antiepileptic drug used in the treatment of patients with IGEs, is thought to primarily provide efficacy by inhibiting GABA transaminase to promote GABAergic transmission and through inhibition of sodium channels; this compound is also known to block T-type channels (reviewed in Czapinski et al., 2005). Initial studies report the partial inhibition of T-type channel currents at

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high concentrations in rat nodose ganglion neurons relative to the therapeutic plasma concentration (Kelly et al., 1990; Todorovic and Lingle, 1998). However, even at cloned Cav3.1 channels, 10 mM valproate only modestly inhibited the Ca2þ channel current by 10% (Lacinova et al., 2000). More recently, the antagonistic action of valproate and ethosuximide on LVA channels of thalamic relay neurons in WAG/Rij rats has been systematically compared to nonepileptic control rats (Broicher et al., 2007b). Both ethosuximide (0.25–0.75 mM) and valproate (1 mM) inhibit T-type Ca2þ channel currents with higher affinity in acutely isolated thalamocortical relay neurons from WAG/Rij rats than in comparable neurons from nonepileptic rats. In addition, ethosuximide delays the onset of the low-threshold spike and increases the tonic action potentials in thalamocortical neurons from WAG/Rij rats, an effect that is different to that observed in control rats. A role for T-type channels in temporal lobe seizures has been suggested as fluoxetine, a reuptake inhibitor of serotonin used to treat depression, not only inhibits LVA (IC50 ¼ 6.8 mM) and HVA (
1–2 mM) calcium channels in cultured hippocampal pyramidal cells, but also reduces Kþ-induced seizure-like activity in hippocampal slices (Wong et al., 1995; De
ak et al., 2000). In addition, in rodents, fluoxetine dose-dependently protects animals from limbic seizures evoked by bicuculline applied into the deep prepiriform cortex (Prendiville and Gale, 1993) as well as by increasing the after-discharge threshold of hippocampal seizures induced by electrical stimulation (Wada et al., 1995). In genetically epilepsy-prone rats (GEPR), fluoxetine in a dose-dependent manner reduces convulsions induced by sound stimulus (Dailey et al., 1996). Interestingly, in humans when fluoxetine is coadministered with valproate, carbamazepine, or phenobarbital or in mice with phenytoin, carbamazepine, and ameltolide, the effects of the AEDS are enhanced (Leander, 1992; Favale et al., 1995). Other studies examining roles of T-type channels in partial and generalized seizures have proposed that the therapeutic action of phenytoin and zonisamide may be in part due to T-type inhibition (Suzuki et al., 1992; Todorovic et al., 2000). Phenytoin is used clinically to treat partial and generalized seizures and is known to primarily inhibit sodium channels. It also inhibits recombinant T-type channels at concentrations close to the maximal therapeutic concentration (Cav3.1 IC50 ¼ 124 mM; Cav3.2 IC50 ¼ 8.3 and 192 mM). Zonisamide, an adjunctive therapy in adults with partial onset seizures with multiple sites of action, shows T-type current blockade in cultured neurons isolated from rat cerebral cortex of 38% at 50 mM (Suzuki et al., 1992). In addition, flunarizine, a potent blocker of recombinant Cav3.1 (IC50 ¼ 0.53 mM), Cav3.2 (IC50 ¼ 0.36 mM), and Cav3.3 (IC50 ¼ 0.84 mM) channels, also exhibits anticonvulsant properties in both animals and humans (Santi et al., 2002). In humans, flunarizine administered as an adjuvant therapy is efficacious in patients with complex partial seizures with or without secondarily generalized seizures as well as reflex seizures (Starreveld et al., 1989; Durrheim et al., 1992; Pledger et al., 1994). In various chemoconvulsant and electroconvulsant animal models, flunarizine monotherapy or coadministration results in protection (Desmedt et al., 1975; De Sarro et al., 1986, 1992; Drago et al., 1986; Pohl and Mares, 1987; Mack and Gilbert, 1992; Rodger and Pleuvry, 1993; Becker and Grecksch, 1995; Joseph et al., 1998a, 1998b), though in some models no protection is

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observed for cocaine-induced seizures, cortical stimulation, amygdala kindling, or pentylenetetrazol seizures (Trommer and Pasternak, 1989; Gasior et al., 1996, 1999).

2.4 CONCLUSIONS In summary, there is considerable pharmacological, genetic, and physiological evidence implicating N-type and T-type calcium channels in a number of pathophysiological conditions centered around neuronal hyperexcitability. Although there exist clinical agents that nonselectively block T-type calcium channels, there are no subtype-specific drugs yet available; thus, there remains both significant clinical challenge and opportunity. In the case of N-type channels and intractable pain, while Prialt provides excellent clinical proof of concept, its peptidic nature requiring intrathecal administration together with its narrow therapeutic window provides the opportunity for developing equally efficacious but safer and orally available N-type channel blockers.

REFERENCES Adams DJ, Smith AB, Schroeder CI, Yasuda T, Lewis RJ, 2003. Omega-conotoxin CVID inhibits a pharmacologically distinct voltage-sensitive calcium channel associated with transmitter release from preganglionic nerve terminals. J Biol Chem 278(6): 4057– 4062. Altier C, Dale CS, Kisilevsky AE, Chapman K, Castiglioni AJ, Matthews EA, Evans RM, Dickenson AH, Lipscombe D, Vergnolle N, Zamponi GW, 2007. Differential role of N-type calcium channel splice isoforms in pain. J Neurosci 27(24): 6363–6373. Babbedge RC, Soper AJ, Gentry CT, Hood VC, Campbell EA, Urban L, 1996. In vitro characterization of a peripheral afferent pathway of the rat after chronic sciatic nerve section. J Neurophysiol 76(5): 3169–3177. Baccei ML, Kocsis JD, 2000. Voltage-gated calcium currents in axotomized adult rat cutaneous afferent neurons. J Neurophysiol 83(4): 2227–2238. Barton ME, Eberle EL, Shannon HE, 2005. The antihyperalgesic effects of the T-type calcium channel blockers ethosuximide, trimethadione, and mibefradil. Eur J Pharmacol 521(1–3): 79–85. Becker A, Grecksch G, 1995. Flunarizine—its effect on pentylenetetrazol-kindled seizures and on related cognitive disturbances. Pharmacol Biochem Behav 52(4): 765–769. Bell TJ, Thaler C, Castiglioni AJ, Helton TD, Lipscombe D, 2004. Cell-specific alternative splicing increases calcium channel current density in the pain pathway. Neuron 41(1): 127–138. Bennett GJ, Xie YK,1988. A peripheral mononeuropathy in rat that produces disorders of pain sensation like those seen in man. Pain 33(1): 87–107. Blake DW, Scott DA, Angus JA, Wright CE, 2005. Synergy between intrathecal omegaconotoxin CVID and dexmedetomidine to attenuate mechanical hypersensitivity in the rat. Eur J Pharmacol 506(3): 221–227.

54

VOLTAGE-GATED N-TYPE AND T-TYPE CALCIUM CHANNELS

Boland LM, Morrill JA, Bean BP, 1994. Omega-conotoxin block of N-type calcium channels in frog and rat sympathetic neurons. J Neurosci 14: 5011–5027. Boucher TJ, Okuse K, Bennett DL, Munson JB, Wood JN, McMahon SB, 2000. Potent analgesic effects of GDNF in neuropathic pain states. Science 290(5489): 124–127. Bourinet E, Alloui A, Monteil A, Barrere C, Couette B, Poirot O, Pages A, McRory J, Snutch TP, Eschalier A, Nargeot J, 2005. Silencing of the Cav3.2 T-type calcium channel gene in sensory neurons demonstrates its major role in nociception. EMBO J 24(2): 315–324. Bowersox SS, Gadbois T, Singh T, Pettus M, Wang YX, Luther RR, 1996. Selective N-type neuronal voltage-sensitive calcium channel blocker, SNX-111, produces spinal antinociception in rat models of acute, persistent and neuropathic pain. J Pharmacol Exp Ther 279: 1243–1249. Broicher T, Kanyshkova T, Landgraf P, Rankovic V, Meuth P, Meuth SG, Pape HC, Budde T, 2007a. Specific expression of low-voltage-activated calcium channel isoforms and splice variants in thalamic local circuit interneurons. Mol Cell Neurosci 36: 132–145. Broicher T, Seidenbecher T, Meuth P, Munsch T, Meuth SG, Kanyshkova T, Pape HC, Budde T, 2007b. T-current related effects of antiepileptic drugs and a Ca2þ channel antagonist on thalamic relay and local circuit interneurons in a rat model of absence epilepsy. Neuropharmacology 53(3): 431–446. Brose WG, Gutlove DP, Luther RR, Bowersox SS, McGuire D, 1997. Use of intrathecal SNX111, a novel, N-type, voltage-sensitive, calcium channel blocker, in the management of intractable brachial plexus avulsion pain. Clin J Pain 13: 256–259. Carbone E, Lux HD, 1984a. A low voltage-activated calcium conductance in embryonic chick sensory neurons. Biophys J 46(3): 413–418. Carbone E, Lux HD, 1984b. A low voltage-activated, fully inactivating Ca channel in vertebrate sensory neurons. Nature 310(5977): 501–502. Chen Y, Lu J, Pan H, Zhang Y, Wu H, Xu K, Liu X, Jiang Y, Bao X, Yao Z, Ding K, Lo WH, Qiang B, Chan P, Shen Y, Wu X, 2003a. Association between genetic variation of CACNA1H and childhood absence epilepsy. Ann Neurol 54(2): 239–243. Chen Y, Lu J, Zhang Y, Pan H, Wu H, Xu K, Liu X, Jiang Y, Bao X, Zhou J, Liu W, Shi G, Shen Y, Wu X, 2003b. T-type calcium channel gene alpha (1G) is not associated with childhood absence epilepsy in the Chinese Han population. Neurosci Lett 341(1): 29–32. Cheng JK, Chen CC, Yang JR, Chiou LC, 2006. The antiallodynic action target of intrathecal gabapentin: Ca2þ channels, KATP channels or N-methyl-D-aspartic acid receptors? Anesth Analg 102(1): 182–187. Cheng JK, Lin CS, Chen CC, Yang JR, Chiou LC, 2007. Effects of intrathecal injection of T-type calcium channel blockers in the rat formalin test. Behav Pharmacol 8(1): 1–8. Chioza B, Everett K, Aschauer H, Brouwer O, Callenbach P, Covanis A, Dulac O, Durner M, Eeg-Olofsson O, Feucht M, Friis M, Heils A, Kjeldsen M, Larsson K, Lehesjoki AE, Nabbout R, Olsson I, Sander T, Siren A, Robinson R, Rees M, Gardiner RM, 2006. Evaluation of CACNA1H in European patients with childhood absence epilepsy. Epilepsy Res 69(2): 177–181. Choi S, Na HS, Kim J, Lee J, Lee S, Kim D, Park J, Chen CC, Campbell KP, Shin HS, 2007. Attenuated pain responses in mice lacking Ca(V)3.2 T-type channels. Genes Brain Behav 6(5): 425–431.

REFERENCES

55

Cizkova D, Marsala J, Lukacova N, Marsala M, Jergova S, Orendacova J, Yaksh TL, 2002. Localization of N-type Ca2þ channels in the rat spinal cord following chronic constrictive nerve injury. Exp Brain Res 147(4): 456–463. Coulter DA, Huguenard JR, Prince DA, 1989a. Calcium currents in rat thalamocortical relay neurones: kinetic properties of the transient low-threshold current J Physiol 414: 587–604. Coulter DA, Huguenard JR, Prince DA, 1989b. Characterization of ethosuximide reduction of low-threshold calcium current in thalamic neurons. Ann Neurol 25(6): 582–593. Craig PJ, Beattie RE, Folly EA, Banerjee MD, Reeves MB, Priestley JV, Carney SL, Sher E, Perez-Reyes E, Volsen SG, 1999. Distribution of the voltage-dependent calcium channel alpha1G subunit mRNA and protein throughout the mature rat brain. Eur J Neurosci 11(8): 2949–2964. Czapinski P, Blaszczyk B, Czuczwar SJ, 2005. Mechanisms of action of antiepileptic drugs. Curr Top Med Chem 5(1): 3–14. Dailey JW, Yan QS, Adams-Curtis LE, Ryu JR, Ko KH, Mishra PK, Jobe PC, 1996. Neurochemical correlates of antiepileptic drugs in the genetically epilepsy-prone rat (GEPR). Life Sci 58(4): 259–266. De
ak F, Laszt
oczi B, Pacher P, Pethe€o GL, Kecskem
eti V, Sp€at A, 2000. Inhibition of voltagegated calcium channels by fluoxetine in rat hippocampal pyramidal cells. Neuropharmacology 39(6): 1029–1036. De la Pena E, Geijo-Barrientos E, 1996. Laminar localization, morphology, and physiological properties of pyramidal neurons that have the low-threshold calcium current in the guineapig medial frontal cortex. J Neurosci 16(17): 5301–5311. De Sarro GB, Nistico G, Meldrum BS, 1986. Anticonvulsant properties of flunarizine on reflex and generalized models of epilepsy. Neuropharmacology 25(7): 695–701. De Sarro GB, De Sarro A, Trimarchi GR, Nistico G, 1992. Effects of some calcium antagonists upon the activity of common antiepileptic compounds on sound-induced seizures in DBA/2 mice. Gen Pharmacol 23(1): 75–82. Deschênes M, Paradis M, Roy JP, Steriade M, 1984. Electrophysiology of neurons of lateral thalamic nuclei in cat: resting properties and burst discharges. J Neurophysiol 51(6): 1196–1219. Desmedt LK, Niemegeers CJ, Janssen PA, 1975. Anticonvulsive properties of cinnarizine and flunarizine in rats and mice. Arzneimittelforschung 25(9): 1408–1413. Destexhe A, Sejnowski TJ, 2002. The initiation of bursts in thalamic neurons and the cortical control of thalamic sensitivity. Philos Trans R Soc Lond B Biol Sci 357(1428): 1649–1657. Devor M, Janig W, Michaelis M, 1994. Modulation of activity in dorsal root ganglion neurons by sympathetic activation in nerve-injured rats. J Neurophysiol 71(1): 38–47. Diaz A, Dickenson AH, 1997. Blockade of spinal N- and P-type but not L-type calcium channels inhibits the excitability of rat dorsal horn neurons produced by subcutaneous formalin inflammation. Pain 69(1–2): 93–100. Dickenson AH, Sullivan AF, 1987. Subcutaneous formalin-induced activity of dorsal horn neurons in the rat: differential response to an intrathecal opiate administered pre or post formalin. Pain 30(3): 349–360. Djouhri L, Koutsikou S, Fang X, McMullan S, Lawson SN, 2006. Spontaneous pain, both neuropathic and inflammatory, is related to frequency of spontaneous firing in intact C-fiber nociceptors. J Neurosci 26(4): 1281–1292.

56

VOLTAGE-GATED N-TYPE AND T-TYPE CALCIUM CHANNELS

Dogrul A, Yeşilyurt O, Işimer A, G€uzeldemir ME, 2001. L-type and T-type calcium channel blockade potentiate the analgesic effects of morphine and selective mu opioid agonist but not to selective delta and kappa agonist at the level of the spinal cord in mice. Pain 93(1): 61–68. Dogrul A, Gardell LR, Ossipov MH, Tulunay FC, Lai J, Porreca F, 2003. Reversal of experimental neuropathic pain by T-type calcium channel blockers. Pain 105(1–2): 159–168. Drago F, Valerio C, Clementi G, Scapagnini U, 1986. Effects of flunarizine on experimentally induced convulsions in animals. Funct Neurol 1(4): 529–532. Drake ME, Greathouse NI, Renner JB, Armentbright AD, 2004. Open-label zonisamide for refractory migraine. Clin Neuropharmacol 27(6): 278–280. Dubel SJ, Starr TV, Hell J, Ahlijanian MK, Enyeart JJ, Catterall WA, Stch TP, 1992. Molecular cloning of the alpha-1 subunit of an omega-conotoxin sensitive calcium channel. Proc Natl Acad Sci USA 89: 5058–5062. Dubreuil AS, Boukhaddaoui H, Desmadryl G, Martinez-Salgado C, Moshourab R, Lewin GR, Carroll P, Valmier J, Scamps F, 2004. Role of T-type calcium current in identified D-hair mechanoreceptor neurons studied in vitro. J Neurosci 24(39): 8480–8484. Durrheim DN, Joubert J, Griesel RD, 1992. Flunarizine—effective add-on therapy in reading epilepsy. S Afr Med J 82(1): 21–23. Ertel SI, Ertel EA, Clozel JP, 1997. T-type Ca2þ channels and pharmacological blockade: potential pathophysiological relevance. Cardiovasc Drugs Ther 11(6): 723–739. Evans AR, Nicol GD, Vasko MR, 1996. Differential regulation of evoked peptide release by voltage-sensitive calcium channels in the rat sensory neurons. Brain Res 712: 265–273. Faas GC, Vreugdenhil M, Wadman WJ, 1996. Calcium currents in pyramidal CA1 neurons in vitro after kindling epileptogenesis in the hippocampus of the rat. Neuroscience 75(1): 57–67. Favale E, Rubino V, Mainardi P, Lunardi G, Albano C, 1995. Anticonvulsant effect of fluoxetine in humans. Neurology 45(10): 1926–1927. Feng ZP, Doering CJ, Winkfein RJ, Beedle AM, Spafford JD, Zamponi GW, 2003. Determinants of inhibition of transiently expressed voltage-gated calcium channels by omegaconotoxins GVIA and MVIIA. J Biol Chem 278: 20171–20178. Finnerup NB, Otto M, McQuay HJ, Jensen TS, Sindrup SH, 2005. Algorithm for neuropathic pain treatment: an evidence-based proposal. Pain 118(3): 289–305. Flatters SJ, Bennett GJ, 2004. Ethosuximide reverses paclitaxel- and vincristine-induced painful peripheral neuropathy. Pain 109(1–2): 150–161. Foong FW, Satoh M, 1983. Analgesic potencies of non-narcotic, narcotic and anesthetic drugs as determined by the bradykinin-induced biting-like responses in rats. Jpn J Pharmacol 33(5): 933–938. Fuchs A, Rigaud M, Sarantopoulos CD, Filip P, Hogan QH, 2007. Contribution of calcium channel subtypes to the intracellular calcium signal in sensory neurons: the effect of injury. Anesthesiology 107(1): 117–127. Fukuizumi T, Ohkubo T, Kitamura K, 2003a. Spinally delivered N-, P/Q- and L-type Ca2þchannel blockers potentiate morphine analgesia in mice. Life Sci 73: 2873–2881. Fukuizumi T, Ohkubo T, Kitamura K, 2003b. Spinal sensitization mechanism in vincristineinduced hyperalgesia in mice. Neurosci Lett 343: 89–92.

REFERENCES

57

Gasior M, Kaminski R, Brudniak T, Kleinrok Z, Czuczwar SJ, 1996. Influence of nicardipine, nimodipine and flunarizine on the anticonvulsant efficacy of antiepileptics against pentylenetetrazol in mice. J Neural Transm 103(7): 819–831. Gasior M, Ungard JT, Witkin JM, 1999. Preclinical evaluation of newly approved and potential antiepileptic drugs against cocaine-induced seizures. J Pharmacol Exp Ther 290(3): 1148–1156. Gohil K, Bell JR, Ramachandran J. Miljanich GP, 1994. Neuroanatomical distribution of receptors for a novel voltage-sensitive calcium channel antagonist, SNX-230 (w-conopeptide MVIIC). Brain Res 653: 258–266. Gomora JC, Daud AN, Weiergraber M, Perez-Reyes E, 2001. Block of cloned human T-type calcium channels by succinimide antiepileptic drugs. Mol Pharmacol 60: 1121–1132. Green MW, Selman JE, 1991. Review article: the medical management of trigeminal neuralgia. Headache 31(9): 588–592. Gruner W. Silva LR, 1994. w-Conotoxin sensitivity and presynaptic inhibition of glutamatergic sensory neurotransmission in vitro. J Neurosci 14: 2800–2808. Guay DR, 2003. Oxcarbazepine, topiramate, zonisamide, and levetiracetam: potential use in neuropathic pain. Am J Geriatr Pharmacother 1(1): 18–37. Harper AA, Lawson SN, 1985a. Conduction velocity is related to morphological cell type in rat dorsal root ganglion neurons. J Physiol 359: 31–46. Harper AA, Lawson SN, 1985b. Electrical properties of rat dorsal root ganglion neurons with different peripheral nerve conduction velocities. J Physiol 359: 47–63. Herlitze S, Garcia DE, Mackie K, Hille B, Scheuer T, Catterall WA, 1996. Modulation of Ca2þ channels by G-protein beta gamma subunits. Nature 380(6571): 258–262. Heron SE, Phillips HA, Mulley JC, Mazarib A, Neufeld MY, Berkovic SF, Scheffer IE, 2004. Genetic variation of CACNA1H in idiopathic generalized epilepsy. Ann Neurol 55: 595–596. Heron SE, Khosravani H, Varela D, Bladen C, Williams TC, Newman MR, Scheffer IE, Berkovic SF, Mulley JC, Zamponi GW, 2007. Extended spectrum of idiopathic generalized epilepsies associated with CACNA1H functional variants. Ann Neurol 62(6): 560–568. Hildebrand ME, Snutch TP, 2006. Contributions of T-type calcium channels to the pathophysiology of pain signaling. Drug Discovery Today: Dis. Mech. 3: 335–341. Hogan QH, 2007. Role of decreased sensory neuron membrane calcium currents in the genesis of neuropathic pain. Croat Med J 48(1): 9–21. Holz GG, Dunlap K, Kream RM, 1988. Characterization of the electrically evoked release of substance P from dorsal root ganglion neurons: methods and dihydropyridine sensitivity. J Neurosci 8: 463–471. Hord AH, Denson DD, Chalfoun AG, Azevedo MI, 2003. The effect of systemic zonisamide (Zonegran) on thermal hyperalgesia and mechanical allodynia in rats with an experimental mononeuropathy. Anesth Analg 96(6): 1700–1706. Huguenard JR, 2002. Block of T-type Ca2þ channels is an important action of succinimide antiabsence drugs. Epilepsy Curr 2(2): 49–52. Huguenard JR, Prince DA, 1992. A novel T-type current underlies prolonged Ca2þ-dependent burst firing in GABAergic neurons of rat thalamic reticular nucleus. J Neurosci 12(10): 3804–3817. Huguenard JR, Prince DA, 1994. Intrathalamic rhythmicity studied in vitro: nominal T-current modulation causes robust antioscillatory effects. J Neurosci 14(9): 5485–5502. Ikeda SR, 1996. Voltage-dependent modulation of N-type calcium channels by G-protein beta gamma subunits. Nature 380(6571): 255–258.

58

VOLTAGE-GATED N-TYPE AND T-TYPE CALCIUM CHANNELS

Ikeda SR, Dunlap K, 2007. Calcium channels diversify their signaling portfolio. Nat Neurosci 10(3): 269–271. Ino M, Yoshinaga T, Wakamori M, Miyamoto N, Takahashi E, Sonoda J, Kagaya T, Oki T, Nagasu T, Nishizawa Y, Tanaka I, Imoto K, Aizawa S, Koch S, Schwartz A, Niidomem T, Sawada K, Mori Y, 2001. Functional disorders of the sympathetic nervous system in mice lacking the a1B subunit (Cav2.2) of N-type calcium channels. Proc Natl Acad Sci USA 98: 5323–5328. Isope P, Murphy TH, 2005. Low threshold calcium currents in rat cerebellar Purkinje cell dendritic spines are mediated by T-type calcium channels. J Physiol 562(Part 1): 257–269. Jagodic MM, Pathirathna S, Nelson MT, Mancuso S, Joksovic PM, Rosenberg ER, Bayliss DA, Jevtovic-Todorovic V, Todorovic SM, 2007. Cell-specific alterations of T-type calcium current in painful diabetic neuropathy enhance excitability of sensory neurons. J Neurosci 27(12): 3305–3316. Jahnsen H, Llinas R, 1984. Ionic basis for the electro-responsiveness and oscillatory properties of guinea-pig thalamic neurones in vitro. J Physiol 349: 227–247. Jim
enez C, Bourinet E, Leuranguer V, Richard S, Snutch TP, Nargeot J, 2000. Determinants of voltage-dependent inactivation affect mibefradil block of calcium channels. Neuropharmacology 39(1): 1–10. Joseph S, David J, Joseph T, 1998a. Additive anticonvulsant effect of flunarizine and sodium valproate on electroshock and chemoshock induced seizures in mice. Indian J Physiol Pharmacol 42(3): 383–388. Joseph S, David J, Joseph T, 1998b. Synergistic effect of flunarizine and sodium valproate on seizure thresholds elicited by cortical stimulation in conscious rats. Indian J Physiol Pharmacol 42(1): 39–49. Jung HY, Staff NP, Spruston N, 2001. Action potential bursting in subicular pyramidal neurons is driven by a calcium tail current. J Neurosci 21(10): 3312–3321. Kajander KC, Bennett GJ, 1992. Onset of a painful peripheral neuropathy in rat: a partial and differential deafferentation and spontaneous discharge in A beta and A delta primary afferent neurons. J Neurophysiol 68(3): 734–744. Kajander KC, Wakisaka S, Bennett GJ, 1992. Spontaneous discharge originates in the dorsal root ganglion at the onset of a painful peripheral neuropathy in the rat. Neurosci Lett 138(2): 225–228. Karst H, Joels M, Wadman WJ, 1993. Low-threshold calcium current in dendrites of the adult rat hippocampus. Neurosci Lett 164(1–2): 154–158. Kato A, Ohkubo T, Kitamura K, 2002. Algogen-specific pain processing in mouse spinal cord: differential involvement of voltage-dependent Ca2þ channels in synaptic transmission. Br J Pharmacol 135(5): 1336–1342. Kelly KM, Gross RA, Macdonald RL, 1990. Valproic acid selectively reduces the lowthreshold (T) calcium current in rat nodose neurons. Neurosci Lett 116(1–2): 233– 238. Kerr LM, Filloux F, Olivera BM, Jackson H, Wamsley JK, 1988. Autoradiographic localization of calcium channels with [125I]omega-conotoxin in rat brain. Eur J Pharmacol 146: 181–183. Khosravani H, Altier C, Simms B, Hamming KS, Snutch TP, Mezeyova J, McRory JE, Zamponi GW, 2004. Gating effects of mutations in the Cav3.2 T-type calcium channel associated with childhood absence epilepsy. J Biol Chem 279(11): 9681–9684.

REFERENCES

59

Khosravani H, Bladen C, Parker DB, Snutch TP, McRory JE, Zamponi GW, 2005. Effects of Cav3.2 channel mutations linked to idiopathic generalized epilepsy. Ann Neurol 57(5): 745–749. Kim C, Jun K, Lee T, Kim S-S, McEnery MW, Chin H, Kim H-L, Park JM, Kim DW, Jung SJ, Kim J, Shin H-S, 2001. Altered nociceptive response in mice deficient in the a1B subunit of the voltage-dependent Ca channel. Mol Cell Neurosci 18: 235–245. Kim D, Song I, Keum S, Lee T, Jeong MJ, Kim SS, McEnery MW, Shin HS, 2001. Lack of the burst firing of thalamocortical relay neurons and resistance to absence seizures in mice lacking alpha(1G) T-type Ca2þ channels. Neuron 31(1): 35–45. Kim D, Park D, Choi S, Lee S, Sun M, Kim C, Shin HS, 2003. Thalamic control of visceral nociception mediated by T-type Ca2þ channels. Science 302(5642): 117–119. Kito M, Maehara M, Watanabe K, 1996. Mechanisms of T-type calcium channel blockade by zonisamide. Seizure 5(2): 115–119. Koyano K, Abe T, Nishiuchi Y, Sakakibara S, 1987. Effects of synthetic omega-conotoxin on synaptic transmission. Eur J Pharmacol 135(3): 337–343. Kyi M, Lopaticki S, O’Brien TJ, Reid C, Foote SJ, Genetic sequence and mRNA expression of two Ca2þ channel genes (Cav3.2 & CACNG2) in genetic absence epilepsy rats from Strasbourg. Australian Neuroscience Society, Online 2006, Abstract Viewer and Itinerary Planner, Program No. ORAL-11-03. Lacinova L, Klugbauer N, Hofmann F, 2000. Regulation of the calcium channel alpha(1G) subunit by divalent cations and organic blockers. Neuropharmacology 39(7): 1254–1266. Leander JD. 1992. Fluoxetine, a selective serotonin-uptake inhibitor, enhances the anticonvulsant effects of phenytoin, carbamazepine, and ameltolide (LY201116). Epilepsia 33(3): 573–576. Lechin F, van der Dijs B, Lechin ME, Amat J, Lechin AE, Cabrera A, Gomez F, Acosta E, Arocha L, Villa S, 1989. Pimozide therapy for trigeminal neuralgia. Arch Neurol 46(9): 960–963. Leresche N, Parri HR, Erdemli G, Guyon A, Turner JP, Williams SR, Asprodini E, Crunelli V, 1998. On the action of the anti-absence drug ethosuximide in the rat and cat thalamus. J Neurosci 18(13): 4842–4853. Lewis RJ, Nielsen KJ, Craik DJ, Loughnan ML, Adams DA, Sharpe IA, Luchian T, Adams DJ, Bond T, Thomas L, Jones A, Matheson JL, Drinkwater R, Andrews PR, Alewood PF, 2000. Novel omega-conotoxins from Conus catus discriminate among neuronal calcium channel subtypes. J Biol Chem 275: 35335–35344. Lia EN, Prado WA, 1999. Effects of intrathecal L- and N-type calcium channel blockers on the antinociception evoked by opioid agonists in the rat tail flick test. Acta Physiol Pharmacol Ther Latinoam 49(4): 195–203. Liang J, Zhang Y, Wang J, Pan H, Wu H, Xu K, Liu X, Jiang Y, Shen Y, Wu X, 2006. New variants in the CACNA1H gene identified in childhood absence epilepsy. Neurosci Lett 406 (1–2): 27–32. Liang J, Zhang Y, Chen Y, Wang J, Pan H, Wu H, Xu K, Liu X, Jiang Y, Shen Y, Wu X, 2007. Common polymorphisms in the CACNA1H gene associated with childhood absence epilepsy in Chinese Han population. Ann Hum Genet 71 (Part 3): 325–335. Liu CN, Wall PD, Ben-Dor E, Michaelis M, Amir R, Devor M, 2000. Tactile allodynia in the absence of C-fiber activation: altered firing properties of DRG neurons following spinal nerve injury. Pain 85(3): 503–521.

60

VOLTAGE-GATED N-TYPE AND T-TYPE CALCIUM CHANNELS

Llin
as R, Jahnsen H, 1982. Electrophysiology of mammalian thalamic neurones in vitro. Nature 297(5865): 406–408. Luo ZD, Chaplan SR, Higuera ES, Sorkin LS, Stauderman KA, Williams ME, Yaksh TL, 2001. Upregulation of dorsal root ganglion (alpha)2(delta) calcium channel subunit and its correlation with allodynia in spinal nerve-injured rats. J Neurosci 21(6): 1868–1875. Mack CM, Gilbert ME, 1992. An examination of the anticonvulsant properties of voltagesensitive calcium channel inhibitors in amygdala kindled seizures. Psychopharmacology (Berl) 106(3): 365–369. Malmberg AB, Yaksh TL, 1994. Voltage-sensitive calcium channels in spinal nociceptive processing: blockade of N-and P-type channels inhibits formalin-induced nociception. J Neurosci 14: 4882–4890. Malmberg AB, Yaksh TL, 1995. Effect of continuous intrathecal infusion of omega- conopeptides, N-type calcium-channel blockers, on behavior and antinociception in the formalin and hot-plate tests in rats. Pain 60(1): 83–90. Magee JC, Johnston D, 1995. Characterization of single voltage-gated Naþ and Ca2þ channels in apical dendrites of rat CA1 pyramidal neurons. J Physiol 487 (Part 1): 67–90. Maggi CA, Tramontana M, Cecconi R, Santicioli P, 1990. Neurochemical evidence for the involvement of N-type calcium channels in transmitter secretion from peripheral endings of sensory nerves in guinea pigs. Neurosci Lett 144: 203–206. Manning JP, Richards DA, Leresche N, Crunelli V, Bowery NG, 2004. Cortical-area specific block of genetically determined absence seizures by ethosuximide. Neuroscience 123(1): 5–9. Markram H, Sakmann B, 1994. Calcium transients in dendrites of neocortical neurons evoked by single subthreshold excitatory postsynaptic potentials via low-voltage-activated calcium channels. Proc Natl Acad Sci USA 91(11): 5207–5211. Martin RL, Lee JH, Cribbs LL, Perez-Reyes E, Hanck DA, 2000. Mibefradil block of cloned Ttype calcium channels. J Pharmacol Exp Ther 295(1): 302–308. Mathur VS, 2000. Ziconotide: a new pharmacological class of drug for the management of pain. Semin Anesth Perioperat Med Pain 19: 67–75. Matsuki N, Quandt FN, Ten Eick RE, Yeh JZ, 1984. Characterization of the block of sodium channels by phenytoin in mouse neuroblastoma cells. J Pharmacol Exp Ther 228(2): 523–530. Matthews EA, Dickenson AH, 2001a. Effects of ethosuximide, a T-type Ca2þ channel blocker, on dorsal horn neuronal responses in rats. Eur J Pharmacol 415(2–3): 141–149. Matthews EA, Dickenson AH, 2001b. Effects of spinally delivered N- and P-type voltagedependent calcium channel antagonists on dorsal horn neuronal responses in a rat model of neuropathy. Pain 92(1–2): 235–246. McCall WD, Tanner KD, Levine JD, 1996. Formalin induces biphasic activity in C-fibers in the rat. Neurosci Lett 208(1): 45–48. McCallum JB, Kwok WM, Mynlieff M, Bosnjak ZJ, Hogan QH, 2003. Loss of T-type calcium current in sensory neurons of rats with neuropathic pain. Anesthesiology 98(1): 209–216. McCleane GJ, 1999. Intravenous infusion of phenytoin relieves neuropathic pain: a randomized, double-blinded, placebo-controlled, crossover study. Anesth Analg 89(4): 985–988. McKay BE, McRory JE, Molineux ML, Hamid J, Snutch TP, Zamponi GW, Turner RW, 2006. Ca(V)3 T-type calcium channel isoforms differentially distribute to somatic and dendritic compartments in rat central neurons. Eur J Neurosci 24(9): 2581–2594.

REFERENCES

61

McLachlan EM, Janig W, Devor M, Michaelis M, 1993. Peripheral nerve injury triggers noradrenergic sprouting within dorsal root ganglia. Nature 363(6429): 543–546. Meeren HK, Pijn JP, Van Luijtelaar EL, Coenen AM, Lopes da Silva FH, 2002. Cortical focus drives widespread corticothalamic networks during spontaneous absence seizures in rats. J Neurosci 22(4): 1480–1495. Mouginot D, Bossu JL, Gahwiler BH, 1997. Low-threshold Ca2þ currents in dendritic recordings from Purkinje cells in rat cerebellar slice cultures. J Neurosci 17(1): 160– 170. Murakami M, Nakagawasai O, Fujii S, Kameyama K, Murakami S, Hozumi S, Esashi A, Taniguchi R, Yanagisawa T, Tan-no K, Tadano T, Kitamura K, Kisara K, 2001. Antinociceptive action of amlodipine blocking N-type Ca2þ channels at the primary afferent neurons in mice. Eur J Pharmacol 419(2–3): 175–181. Murakami M, Nakagawasai O, Suzuki T, Mobarakeh II, Sakurada Y, Murata A, Yamadera F, Miyoshi I, Yanai K, Tan-No K, Sasano H, Tadano T, Iijima T, 2004. Antinociceptive effect of different types of calcium channels inhibitors and the distribution of the various calcium channel a1 subunits in the dorsal horn of spinal cord in mice. Brain Res 1024: 122–129. Nahm SS, Jung KY, Enger MK, Griffith WH, Abbott LC, 2005. Differential expression of T-type calcium channels in P/Q-type calcium channel mutant mice with ataxia and absence epilepsy. J Neurobiol 62(3): 352–360. Nebe J, Vanegas H, Schaible H, 1998. Spinal application of w-conotoxin-GVIA, an N-type calcium channel antagonist, attenuates enhancement of dorsal spinal neuronal responses caused by intra-articular injection of mustard oil in the rat. Exp Brain Res 120: 61–69. Nelson MT, Joksovic PM, Perez-Reyes E, Todorovic SM, 2005. The endogenous redox agent Lcysteine induces T-type Ca2þ channel-dependent sensitization of a novel subpopulation of rat peripheral nociceptors. J Neurosci 25(38): 8766–8775. Neugebauer V, Vanegas H, Nebe J, R€umenapp P, Schaible HG, 1996. Effects of N- and L-type calcium channel antagonists on the responses of nociceptive spinal cord neurons to mechanical stimulation of the normal and inflamed knee joint. J Neurophysiol 76(6): 3740–3749. Niedermeyer E, 1996. Primary (idiopathic) generalized epilepsy and underlying mechanisms. Clin Electroencephalogr 27(1): 1–21. Nielsen KJ, Schroeder T, Lewis R, 2000. Structure–activity relationships of omega-conotoxins at N-type voltage-sensitive calcium channels. J Mol Recognit 13: 55–70. Nowycky MC, Fox AP, Tsien RW, 1985. Three types of neuronal calcium channel with different calcium agonist sensitivity. Nature 316: 440–443. Olivera BM, McIntosh JM, Cruz LJ, Luque FA, Gray WR, 1984. Purification and sequence of a presynaptic peptide toxin from Conus geographus venom. Biochemistry 23: 5087–5090. Olivera BM, Gray WR, Zeikus R, McIntosh JM, Varga J, Rivier J, de Santos V, Cruz LJ, 1985. Peptide neurotoxins from fish-hunting cone snails. Science 230: 1338–1343. Olivera BM, Cruz LJ, de Santos V, LeCheminant GW, Griffin D, Zeikus R, McIntosh JM, Galyean R, Varga J, Gray WR, 1987. Neuronal calcium channel antagonists. Discrimination between calcium channel subtypes using omega-conotoxin from Conus magus venom. Biochemistry 26: 2086–2090. Omote K, Kawamata M, Satoh O, Iwasaki H, Namiki A, 1996. Spinal antinociceptive action of an N-type voltage-dependent calcium channel blocker on the synergistic interaction with morphine. Anesthesiology 84(3): 636–643.

62

VOLTAGE-GATED N-TYPE AND T-TYPE CALCIUM CHANNELS

Pathirathna S, Brimelow BC, Jagodic MM, Krishnan K, Jiang X, Zorumski CF, Mennerick S, Covey DF, Todorovic SM, Jevtovic-Todorovic V, 2005a. New evidence that both T-type calcium channels and GABAA channels are responsible for the potent peripheral analgesic effects of 5alpha-reduced neuroactive steroids. Pain 114(3): 429–443. Pathirathna S, Todorovic SM, Covey DF, Jevtovic-Todorovic V, 2005b 5Alpha-reduced neuroactive steroids alleviate thermal and mechanical hyperalgesia in rats with neuropathic pain. Pain 117(3): 326–339. Pathirathna S, Covey DF, Todorovic SM, Jevtovic-Todorovic V, 2006. Differential effects of endogenous cysteine analogs on peripheral thermal nociception in intact rats. Pain 125(1–2): 53–64. Peloquin JB, Khosravani H, Barr W, Bladen C, Evans R, Mezeyova J, Parker D, Snutch TP, McRory JE, Zamponi GW, 2006. Functional analysis of Cav3.2 T-type calcium channel mutations linked to childhood absence epilepsy. Epilepsia 47(3): 655–658. Penn RD, Paice JA, 2000. Adverse effects associated with the intrathecal administration of ziconotide. Pain 85(1–2): 291–296. Perez-Reyes E, 2003. Molecular physiology of low-voltage-activated T-type calcium channels. Physiol Rev 83(1): 117–161. Petersen M, Zhang J, Zhang JM, LaMotte RH, 1996. Abnormal spontaneous activity and responses to norepinephrine in dissociated dorsal root ganglion cells after chronic nerve constriction. Pain 67(2–3): 391–397. Pfrieger FW, Veselovsky NS, Gottmann K, Lux HD, 1992. Pharmacological characterization of calcium currents and synaptic transmission between thalamic neurons in vitro. J Neurosci 12 (11): 4347–4357. Pledger GW, Sackellares JC, Treiman DM, Pellock JM, Wright FS, Mikati M, Sahlroot JT, Tsay JY, Drake ME, Olson L, 1994. Flunarizine for treatment of partial seizures: results of a concentration-controlled trial. Neurology 44(10): 1830–1836. Pohl M, Mares P, 1987. Effects of flunarizine on metrazol-induced seizures in developing rats. Epilepsy Res 1(5): 302–305. Polack PO, Guillemain I, Hu E, Deransart C, Depaulis A, Charpier S, 2007. Deep layer somatosensory cortical neurons initiate spike-and-wave discharges in a genetic model of absence seizures. J Neurosci 27(24): 6590–6599. Prendiville S, Gale K, 1993. Anticonvulsant effect of fluoxetine on focally evoked limbic motor seizures in rats. Epilepsia 34(2): 381–384. Raingo J, Castiglioni AJ Lipscombe D, 2007. Alternative splicing controls G proteindependent inhibition of N-type calcium channels in nociceptors. Nat Neurosci 10(3): 285–292. Ramer MS, French GD, Bisby MA, 1997. Wallerian degeneration is required for both neuropathic pain and sympathetic sprouting into the DRG. Pain 72(1–2): 71–78. Richards DA, Manning JP, Barnes D, Rombola L, Bowery NG, Caccia S, Leresche N, Crunelli V, 2003. Targeting thalamic nuclei is not sufficient for the full anti-absence action of ethosuximide in a rat model of absence epilepsy. Epilepsy Res 54(2–3): 97–107. Rodger C, Pleuvry BJ, 1993. Protective effect of flunarizine and nifedipine alone and in combination with anticonvulsant drugs against PTZ-induced seizures in mice. Neuropharmacology 32(3): 257–263. Rojas-Piloni G, Dickenson AH, Condes-Lara M, 2007. Superficial dorsal horn neurons with double spike activity in the rat. Neurosci Lett 419(2): 147–152.

REFERENCES

63

Saddi G, Abbott FV, 2000. The formalin test in the mouse: a parametric analysis of scoring properties. Pain 89(1): 53–63. Saegusa H, Kurihara T, Zong S, Kazuno A, Matsuda Y, Nonaka T, Han W, Toriyama H, Tanabe T, 2001. Suppression of inflammatory and neuropathic pain symptoms in mice lacking the N-type Ca channel. EMBO J 20: 2349–2356. Saegusa H, Matsuda Y, Tanabe T, 2002. Effects of ablation of N- and R-type Ca2þ channels on pain transmission. Neurosci Res 43: 1–7. Sakaue A, Honda M, Tanabe M, Ono H, 2004. Antinociceptive effects of sodium channelblocking agents on acute pain in mice. J Pharmacol Sci 95(2): 181–188. Santi CM, Cayabyab FS, Sutton KG, McRory JE, Mezeyova J, Hamming KS, Parker D, Stea A, Snutch TP, 2002. Differential inhibition of T-type calcium channels by neuroleptics. J Neurosci 22(2): 396–403. Santicioli P, Del Bianco E, Tramontana M, Geppetti P, Maggi CA, 1992. Release of calcitonin gene-related peptide like immunoreactivity induced by electrical stimulation from rat spinal afferents is mediated by conotoxin-sensitive calcium channels. Neurosci Lett 136: 161–164. Schroeder JE, Fischbach PS, McCleskey EW, 1990. T-type calcium channels: heterogeneous expression in rat sensory neurons and selective modulation by phorbol esters. J Neurosci 10(3): 947–951. Scott DA, Wright CE, Angus JA, 2002. Actions of intrathecal w-conotoxins CVIA, GVIA, MVIIA, and morphine in acute and neuropathic pain in the rat. Eur J Pharm 451: 279–286. Scroggs RS, Fox AP, 1992. Calcium current variation between acutely isolated adult rat dorsal root ganglion neurons of different size. J Physiol 445: 639–658. Sherwin AL, 1989. Ethosuximide: clinical use. In: Levy, RH, Mattson, R., Meldrum, B, Penry, JK, Drisfuss, FE, (Eds), Antiepileptic Drugs. Raven Press, New York, pp. 685–689. Shin JB, Martinez-Salgado C, Heppenstall PA, Lewin GR, 2003. A T-type calcium channel required for normal function of a mammalian mechanoreceptor. Nat Neurosci 6(7): 724–730. Silva LR, Amitai Y, Connors BW, 1991. Intrinsic oscillations of neocortex generated by layer 5 pyramidal neurons. Science 251(4992): 432–435. Singh B, Monteil A, Bidaud I, Sugimoto Y, Suzuki T, Hamano S, Oguni H, Osawa M, Alonso ME, Delgado-Escueta AV, Inoue Y, Yasui-Furukori N, Kaneko S, Lory P, Yamakawa K, 2007. Mutational analysis of CACNA1G in idiopathic generalized epilepsy. Hum Mutat 28 (5): 524–525 (Mutation in brief #962 Online). Sluka KA, 1997. Blockade of calcium channels can prevent the onset of secondary hyperalgesia and allodynia induced by intradermal injection of capsaicin in rats. Pain 71: 157–164. Smith MT, Cabot PJ, Ross FB, Robertson AD, Lewis RJ, 2002. The novel N-type calcium channel blocker, AM336, produces potent dose-dependent antinociception after intrathecal dosing in rats and inhibits substance P release in rat spinal cord slices. Pain 96(1–2): 119–127. Snutch TP, 2008. Voltage-gated calcium channels. The New Encyclopedia of Neuroscience, edited by L. Squire et al. Song XJ, Hu SJ, Greenquist KW, Zhang JM, LaMotte RH. 1999. Mechanical and thermal hyperalgesia and ectopic neuronal discharge after chronic compression of dorsal root ganglia. J Neurophysiol 82(6): 3347–3358. Song I, Kim D, Choi S, Sun M, Kim Y, Shin HS, 2004. Role of the alpha1G T-type calcium channel in spontaneous absence seizures in mutant mice. J Neurosci 24(22): 5249–5257.

64

VOLTAGE-GATED N-TYPE AND T-TYPE CALCIUM CHANNELS

Staats PS, Yearwood T, Charapata SG, Presley RW, Wallace MS, Byas-Smith M, Fisher R, Bryce D, Mangieri EA, Luther RR, Mayo M, McGuire D, Ellis D, 2004. Intrathecal ziconotide in the treatment of refractory pain in patients with cancer or AIDS: a randomized controlled trial. J Am Med Assoc 291: 63–70. Starreveld E, de Beukelaar F, Wilson AF, McLean DR, Findlay HP. 1989. Double-blind crossover placebo controlled study of flunarizine in patients with therapy resistant epilepsy. Can J Neurol Sci 16(2): 187–190. Steriade M, Contreras D, 1998. Spike-wave complexes and fast components of cortically generated seizures. I. Role of neocortex and thalamus. J Neurophysiol 80(3): 1439–1455. Su H, Sochivko D, Becker A, Chen J, Jiang Y, Yaari Y, Beck H, 2002. Upregulation of a T-type Ca2þ channel causes a long-lasting modification of neuronal firing mode after status epilepticus. J Neurosci 22(9): 3645–3655. Suh HW, Song DK, Choi SR, Huh SO, Kim YH, 1997. Differential effects of w-conotoxin GVIA, nimodipine, calmidazolium and KN-62 injected intrathecally on the antinociception induced by b-endorphin, morphine and [D-Ala2,N-MePhe4,Gly-ol5]-enkephalin administered intracerebroventrically in the mouse. J Pharmacol Exp Ther 282: 961–966. Suzuki S, Kawakami K, Nishimura S, Watanabe Y, Yagi K, Seino M, Miyamoto K, 1992. Zonisamide blocks T-type calcium channel in cultured neurons of rat cerebral cortex. Epilepsy Res 12(1): 21–27. Takahashi Y, Hashimoto K, Tsuji S, 2004. Successful use of zonisamide for central poststroke pain. J Pain 5(3): 192–194. Talley EM, Cribbs LL, Lee JH, Daud A, Perez-Reyes E, Bayliss DA, 1999. Differential distribution of three members of a gene family encoding low voltage-activated (T-type) calcium channels. J Neurosci 19(6): 1895–1911. Talley EM, Sol
orzano G, Depaulis A, Perez-Reyes E, Bayliss DA, 2000. Low-voltage-activated calcium channel subunit expression in a genetic model of absence epilepsy in the rat. Brain Res Mol Brain Res 75(1): 159–165. Todorovic SM, Lingle CJ, 1998. Pharmacological properties of T-type Ca2þ current in adult rat sensory neurons: effects of anticonvulsant and anesthetic agents. J Neurophysiol 79(1): 240–252. Todorovic SM, Prakriya M, Nakashima YM, Nilsson KR, Han M, Zorumski CF, Covey DF, Lingle CJ, 1998. Enantioselective blockade of T-type Ca2þ current in adult rat sensory neurons by a steroid that lacks gamma-aminobutyric acid-modulatory activity. Mol Pharmacol 54(5): 918–927. Todorovic SM, Perez-Reyes E, Lingle CJ, 2000. Anticonvulsants but not general anesthetics have differential blocking effects on different T-type current variants. Mol Pharmacol 58(1): 98–108. Todorovic SM, Jevtovic-Todorovic V, Meyenburg A, Mennerick S, Perez-Reyes E, Romano C, Olney JW, Zorumski CF, 2001. Redox modulation of T-type calcium channels in rat peripheral nociceptors. Neuron 31(1): 75–85. Todorovic SM, Meyenburg A, Jevtovic-Todorovic V, 2004. Redox modulation of peripheral Ttype Ca2þ channels in vivo: alteration of nerve injury-induced thermal hyperalgesia. Pain 109(3): 328–339. Trommer BL, Pasternak JF, 1989. Flunarizine does not delay the development of generalized seizures by amygdala kindling. Epilepsy Res 4(3): 230–233. Tsakiridou E, Bertollini L, de Curtis M, Avanzini G, Pape HC, 1995. Selective increase in T-type calcium conductance of reticular thalamic neurons in a rat model of absence epilepsy. J Neurosci 15(4): 3110–3117.

REFERENCES

65

Twombly DA, Yoshii M, Narahashi T, 1988. Mechanisms of calcium channel block by phenytoin. J Pharmacol Exp Ther 246(1): 189–195. Urban MO, Ren K, Sablad M, Park KT. 2005. Medullary N-type and P/Q-type calcium channels contribute to neuropathy-induced allodynia. Neuroreport 16(6): 563–566. Vitko I, Chen Y, Arias JM, Shen Y, Wu XR, Perez-Reyes E, 2005. Functional characterization and neuronal modeling of the effects of childhood absence epilepsy variants of CACNA1H, a T-type calcium channel. J Neurosci 25(19): 4844–4855. Vitko I, Bidaud I, Arias JM, Mezghrani A, Lory P, Perez-Reyes E, 2007. The I-II loop controls plasma membrane expression and gating of Ca(v)3.2 T-type Ca2þ channels: a paradigm for childhood absence epilepsy mutations. J Neurosci 27(2): 322–330. Wada Y, Shiraishi J, Nakamura M, Hasegawa H, 1995. Prolonged but not acute fluoxetine administration produces its inhibitory effect on hippocampal seizures in rats. Psychopharmacology (Berl) 118(3): 305–309. Wagner JA, Snowman AM, Biswas A, Olivera BM, Snyder SH, 1988. Omega-conotoxin GVIA binding to a high-affinity receptor in brain: characterization, calcium sensitivity, and solubilization. J Neurosci 8: 3354–3359. Wall PD, Devor M, 1983. Sensory afferent impulses originate from dorsal root ganglia as well as from the periphery in normal and nerve injured rats. Pain 17(4): 321–339. Wallace MS, 2006. Ziconotide: a new nonopioid intrathecal analgesic for the treatment of chronic pain. Expert Rev Neurother 6(10): 423–428. Wang YX, Bezprozvannaya S, Bowersox SS, Nadasdi L, Miljanich G, Mezo G, Silva D, Tarczy-Hornoch K, Luther RR, 1998. Peripheral versus central potencies of N-type voltage-sensitive calcium channel blockers. Naunyn Schmiedebergs Arch Pharmacol 357: 159–168. Wang J, Zhang Y, Liang J, Pan H, Wu H, Xu K, Liu X, Jiang Y, Shen Y, Wu X, 2006. CACNA1I is not associated with childhood absence epilepsy in the Chinese Han population. Pediatr Neurol 35(3): 187–190. Westenbroek RE, Hell JW, Warner C, Dubel SJ, Snutch TP, Catterall WA, 1992. Biochemical properties and subcellular distribution of an N-type calcium channel alpha 1 subunit. Neuron 9: 1099–1115. White G, Lovinger DM, Weight FF, 1989. Transient low-threshold Ca2þ current triggers burst firing through an afterdepolarizing potential in an adult mammalian neuron. Proc Natl Acad Sci USA 86(17): 6802–6806. Williams DS, 1950. A study of thalamic and cortical rhythms in petit mal. Brain 67: 50–69. Winter J, Brevan S, Campbell EA. 1995. Capsaicin and pain mechanisms. Br J Anaesth 75: 157– 168. Wong DT, Bymaster FP, Engleman EA, 1995. Prozac (fluoxetine, Lilly 110140), the first selective serotonin uptake inhibitor and an antidepressant drug: twenty years since its first publication. Life Sci 57(5): 411–441. Wu G, Ringkamp M, Hartke TV, Murinson BB, Campbell JN, Griffin JW, Meyer RA, 2001. Early onset of spontaneous activity in uninjured C-fiber nociceptors after injury to neighboring nerve fibers. J Neurosci 21(8): RC140. Xiao WH, Bennett GJ, 1995. Synthetic omega-conopeptides applied to the site of nerve injury suppress neuropathic pains in rats. J Pharmacol Exp Ther 274(2): 666– 672.

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Yamamoto T, Sakashita Y, 1998. Differential effects of intrathecally administered N- and P-type voltage-sensitive calcium channel blockers upon two models of experimental mononeuropathy in the rat. Brain Res 794: 329–332. Yokoyama K, Kurihara T, Makita K, Tanabe T, 2003. Plastic change in N-type Ca channel expression after preconditioning is responsible for prostaglandin E2-induced long-lasting allodynia. Anesthesiology 99(6): 1364–1370. Zhang J M. Song XJ, LaMotte RH, 1999. Enhanced excitability of sensory neurons in rats with cutaneous hyperalgesia produced by chronic compression of the dorsal root ganglion. J Neurophysiol 82(6): 3359–3366. Zhang Y, Mori M, Burgess DL, Noebels JL, 2002. Mutations in high-voltage-activated calcium channel genes stimulate low-voltage-activated currents in mouse thalamic relay neurons. J Neurosci 22(15): 6362–6371.

3 VOLTAGE-GATED SODIUM CHANNELS: MULTIPLE ROLES IN THE PATHOPHYSIOLOGY OF PAIN SULAYMAN D. DIB-HAJJ1,2, BRYAN C. HAINS1,2, JOEL A. BLACK1,2, 1,2 AND STEPHEN G. WAXMAN 1

Department of Neurology and Center for Neuroscience and Regeneration Research, Yale University School of Medicine, New Haven, CT 06510, USA 2 Rehabilitation Research Center, VA Connecticut Healthcare System, West Haven, CT 06516, USA

3.1 INTRODUCTION Voltage-gated sodium channels underlie the generation and conduction of action potentials (AP) in excitable cells. Sodium channels in the plasma membrane are closed at rest but undergo structural changes in response to depolarization of the cell membrane, leading to cycling of the channel through closed, open, inactive, and repriming states (Hille, 2001). Channel opening is transient, allowing the flow of sodium ions down their concentration gradient, thus generating an inward current, and most channels rapidly inactivate, within milliseconds of opening, and then undergo conformational changes to recover (reprime) from inactivation. The different sodium channel isoforms cycle through these states with different kinetics and voltagedependent properties so that it is possible to identify individual currents produced by some channels in the presence of other members of the family. Sodium channels are heteromultimers of a large a subunit and smaller auxiliary b subunits (Catterall, 2000). The a subunit alone is necessary and sufficient to produce a Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

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functional channel, whereas b subunits and an array of other cytosolic channel partners regulate trafficking and anchoring of the channels at the cell membrane. Still other channel partners modulate biophysical properties of the channels either by direct binding or by inducing posttranslational modifications, for example, phosphorylation (Wood et al., 2004; Abriel and Kass, 2005; Liu et al., 2005; Cantrell, 2001). The a subunits of sodium channels are large polypeptides (size range: 1700–2000 amino acids) that are organized into four domains (DI–DIV), each consisting of six transmembrane segments that are connected by intra- and extracellular linkers (Fig. 3.1) (Catterall, 2000). The b subunits, however, are type I membrane proteins, each with a single transmembrane segment and a larger extracellular domain that has an immunoglobulin fold (Catterall, 2000). Nine a subunits (Nav1.1–Nav1.9) and several of their cognates have been identified in mammals and lower vertebrates (Goldin et al., 2000; Goldin, 2002; Catterall et al., 2005). Many of these channels produce sodium currents with distinct biophysical and pharmacological properties (Rush et al., 2007). Expression of sodium channels is regulated in spatial and temporal patterns, which will be discussed in the coming sections. Because neuropathic pain has peripheral and central components, sodium channel dysregulation in both PNS and CNS may play a role in establishment and maintenance of this pain state. The following sections will review our current knowledge of neuronal sodium channels Nav1.1–1.3, Nav1.6–1.9, with a major focus on those expressed within dorsal root ganglion (DRG) and higher order neurons along pain signaling pathways, and their demonstrable or putative roles in neuropathic pain.

FIGURE 3.1 Schematic of the pore-forming a subunit of voltage-gated sodium channel. The pore-forming subunit of sodium channels is a long polypeptide with 24 transmembrane segments that are organized into four homologous domains (DI–DIV). The N- and C-termini of the channel, and loops 1–3 (L1–L3) that joins the four domains, are cytosolic and have been shown to house sequence motifs for channel partner binding and for phosphorylation of the channel. The binding of different classes of cytosolic proteins and phosphorylation of the channels have been shown to regulate channel trafficking and polarized distribution within neuronal compartments and/or biophysical properties of the channel. The S4 transmembrane segment in each of the domains is a voltage sensor, and the gray sphere in L3 designates the tetrapeptide IFMT, which acts as the fast-inactivating particle of the channel. The extracellular linkers may be sites of N-glycosylation of channels.

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3.2 TISSUE DISTRIBUTION AND SUBCELLULAR LOCALIZATION OF SODIUM CHANNELS Sodium channel expression is tissue and developmentally regulated across the different species where it has been investigated. Neuronal sodium channels, Nav1.1–1.3 and Nav1.6–1.9, can be found within CNS and PNS neurons and within glia, whereas Nav1.4 and Nav1.5 are expressed primarily in skeletal and cardiac myocytes, respectively, (Catterall et al., 2005) and will not be discussed further in this chapter. Most neurons express more than one sodium channel isoform, and the specific channel complement influences the electrogenic properties of these neurons. For example, neurons in adult rodent DRG can express up to five sodium channel isoforms (Fig. 3.2A), and the current profile and voltage-dependent gating properties of the two channels that are resistant to micromolar concentrations of tetrodotoxin can be distinguished even in the presence of other sodium currents (Fig. 3.2B). Dysregulation of sodium channel expression under pathological conditions can lead to neuronal hyper- or hypoexcitability, which contributes to the disease phenotype. The genes for sodium channels Nav1.1, 1.2, 1.3, and 1.7 are clustered on human and mouse chromosome 2 and are thus thought to have a common lineage (Goldin, 2002). Despite their common ancestry and chromosomal location, the expression of these channels is differentially regulated in CNS and PNS tissues and during development. Nav1.1 is expressed in both CNS (Beckh et al., 1989) and DRG neurons (Black et al., 1996) and in cardiac muscle (Maier et al., 2002). Nav1.1 and Nav1.2 are expressed at low levels during embryogenesis and reach adult levels by P14–P21 in the rat CNS (Beckh et al., 1989; Westenbroek et al., 1989; Felts et al., 1997). Nav1.2 is expressed at low levels within DRG neurons during embryogenesis and its expression is significantly attenuated by P0 (Waxman et al., 1994; Black et al., 1996; Schafer et al., 2006). In rodents, Nav1.3 is the predominant isoform during embryogenesis in CNS and DRG neurons; its expression is significantly attenuated after birth (Beckh et al., 1989; Waxman et al., 1994), but adult human CNS neurons continue to produce higher levels of Nav1.3 compared to their rodent counterparts (Whitaker et al., 2001). Nav1.3 transcripts, however, continue to be detected in sympathetic ganglia in adult rodents, at levels comparable to those of other channels (Rush et al., 2006). Nav1.1 and Nav1.3 appear to be predominantly localized to somatodendritic compartments of myelinated neurons (Westenbroek et al., 1989; Whitaker et al., 2001). Recently, Nav1.1 was detected at the initial segment of axons in the inner plexiform layer of the retina (Van Wart et al., 2005); however, Nav1.1 is not detectable in optic nerve axons (Craner et al., 2003). The subcellular distribution of Nav1.1 in adult rat DRG neurons is not well defined. In contrast, Nav1.2 is present along premyelinated axons and is targeted to immature nodes of Ranvier during myelination of CNS axons (Boiko et al., 2001; Kaplan et al., 2001) but is then restricted to the somatodendritic compartment, stretches of axons that lack myelin, and totally nonmyelinated axons (Westenbroek et al., 1989, 1992; Boiko et al., 2001). Considering the very low levels of Nav1.2 in rodent DRG neurons after birth (Waxman et al., 1994; Black et al., 1996), it is not surprising that this channel is not detected at nodes of Ranvier in peripheral myelinated axons (Schafer et al., 2006).

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FIGURE 3.2 Multiple sodium channels and currents in adult DRG neurons. (A) Sodium channel a subunit mRNAs (left panels) and protein (right panels) visualized by subtype-specific riboprobes and antibodies. Transcripts and protein for five different sodium channels (Nav1.1, Nav1.6, Nav1.7, Nav1.8, and Nav1.9) are present at moderate-to-high levels in DRG neurons. Nav1.2 and Nav1.3 are not detectable in adult DRG neurons. Scale bar: 50 mm. (B) Voltagegated sodium currents recorded by whole-cell patch clamp in adult DRG neurons. (a) Only fast, TTX-sensitive sodium current (presumably composed of Nav1.1, Nav1.6, and Nav1.7) is observed in a muscle afferent DRG neuron, which exhibits little overlap between activation (filled circles) and steady-state inactivation (unfilled circles). (b) A small DRG neuron displays only slow, TTX-resistant sodium current (Nav1.8); activation and steady-state inactivation curves are depolarized compared to fast, TTX-sensitive current. (c) Persistent, TTX-resistant sodium current (Nav1.9) recorded from a small DRG neuron from Nav1.8-null mouse. Activation (unfilled circles) and steady-state inactivation (filled circles) show significant overlap (window currents). (Modified and reproduced with permission from Black et al. (1996), Black et al. (2004), Cummins et al. (1999), Sleeper et al. (2000), and Waxman et al. (1999).)

Nav1.6 is located on human chromosome 12 and is expressed in CNS neurons and glia (Burgess et al., 1995; Schaller et al., 1995) and in peripheral sensory neurons (Black et al., 1996; Dietrich et al., 1998) and sympathetic ganglia (Rush et al., 2006). The expression of Nav1.6 in CNS neurons increases after birth, reaching adult levels by

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P14 (Schaller and Caldwell, 2000), whereas it is detected within DRG neurons at E17 and becomes robust by P0 (Felts et al., 1997; Chung, ). Nav1.6 channels have also been detected in immune cells, for example, microglia and macrophages (Craner et al., 2005). The distribution of Nav1.6 in human CNS tissue is in close agreement with that in rodents (Whitaker et al., 1999). Nav1.6 is the major sodium channel at mature nodes of Ranvier in myelinated axons in the CNS, where it replaces Nav1.2 (Boiko et al., 2001; Kaplan et al., 2001) upon the formation of compact myelin and node maturation, and in the PNS (Schafer et al., 2006). In contrast, Nav1.6 is present in the somata and diffusely along substantial lengths of unmyelinated fibers arising from small DRG neurons (Black and Waxman, 2002; Rush et al., 2005). The focal distribution of Nav1.6 at nodes is disrupted in demyelinated lesions in patients with multiple sclerosis (MS) and in mice with experimental autoimmune encephalomyelitis (EAE) (Waxman et al., 2004). The remyelination of experimental lesions with Schwann cells (Black et al., 2006) or olfactory ensheathing cells (Sasaki et al., 2006) leads to the restoration of Nav1.6 immunolabeling at nodes of Ranvier. Nav1.7 is expressed only in sensory and sympathetic neurons in the PNS (Black et al., 1996; Felts et al., 1997; Sangameswaran et al., 1997; Toledo-Aral et al., 1997; Rush et al., 2006) even though it is encoded by the gene SCN9A, which is clustered at the same chromosomal region with SCN1A–SCN3A, the genes that encode Nav1.1–1.3. More important, Nav1.7 is produced by the majority of functionally identified nociceptive neurons (Djouhri et al., 2003). An earlier report of Nav1.7 detection in brain samples (Sangameswaran et al., 1997) has not been confirmed by other studies (Black et al., 1996; Felts et al., 1997; Toledo-Aral et al., 1997). Nav1.7 has also been detected in a variety of other cell types including neuroendocrine cells (Klugbauer et al., 1995; Toledo-Aral et al., 1997), smooth muscle cells (Holm et al., 2002; Jo et al., 2004; Saleh et al., 2005), and in metastatic cancer cells (Diss et al., 2005; Fraser et al., 2005). Nav1.7 is uniformly distributed in the somata and along the unmyelinated Cfibers within the sciatic nerve (Rush et al., 2005). This channel accumulates within neurite tips in DRG neurons in culture (Toledo-Aral et al., 1997), which suggests an analogous accumulation at nerve terminals in vivo. The presence of Nav1.7 at nerve endings would be consistent with the role of this channel in amplifying generation potentials due to its enabling biophysical properties (Cummins et al., 1998). Nav1.8 is a sensory neuron-specific channel that is normally expressed within DRG and trigeminal ganglia but not in CNS neurons (Akopian et al., 1996; Sangameswaran et al., 1996). Nav1.8 is present in most peptidergic and nonpeptidergic small DRG neurons (Fjell et al., 1999; Amaya et al., 2000; Sleeper et al., 2000; Djouhri et al., 2003a; Fang et al., 2005; Rush et al., 2005). Nav1.8 immunostaining signal is abundant in the somata of small DRG neurons (Amaya et al., 2000; Sleeper et al., 2000) and in unmyelinated fibers in the sciatic nerve (Rush et al., 2005). Nav1.8 is also expressed in trigeminal neurons (Bongenhielm et al., 2000; Eriksson et al., 2005) and has been detected in unmyelinated fibers innervating cornea (Black and Waxman, 2002). Nav1.8 is also present within a subpopulation of medium-sized DRG neurons that produce Ab fibers (Rizzo et al., 1994, 1995; Amaya et al., 2000; Djouhri et al., 2003a) and has been reported at
20% of nodes of Ranvier in myelinated axons that innervate tooth pulp (Henry et al., 2005). The reporting of Nav1.8 at nodes of Ranvier in spinal cord tracts

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(Arroyo et al., 2002) is inconsistent with the lack of Nav1.8 message in normal CNS tissue, although a small number of myelinated axons that emanate from DRG neurons and project to the spinal cord might express it. The slowly inactivating Nav1.8 current is absent from proprioceptive, muscle afferent DRG neurons (Rizzo et al., 1994) that have continuous central projections within spinal cord tracts; thus, these cells could not be the source of the putative Nav1.8 signal in that tissue. Nav1.9 is the other sensory neuron-specific sodium channel within DRG and trigeminal ganglia (Dib-Hajj et al., 1998; Tate et al., 1998), and within myenteric neurons (Rugiero et al., 2003). Within DRG neurons, Nav1.9 expression has been reported in small-diameter nonpeptidergic IB4þ neurons (Fjell et al., 1999a; Amaya et al., 2000; Fang et al., 2002; Rush et al., 2005). Low levels of Nav1.9 transcripts were detected by reverse transcription-polymerase chain reaction (RT-PCR) assays in cerebral hemispheres and retina but not in cerebellum or spinal cord, or in satellite or Schwann cells within DRG (Dib-Hajj et al., 1998). Reports of wider distribution of Nav1.9 within CNS tissue (Jeong et al., 2000) have not been confirmed by other studies and are inconsistent with the absence of a persistent TTX-R current, which is attributable to Nav1.9. A report contending that BDNF could activate Nav1.9 channels in CNS neurons (Blum et al., 2002) has yet to be independently reproduced, casting doubt on the validity of the initial observations. Nav1.9 is abundant in the somata of the majority of small nonpeptidergic DRG neurons (Amaya et al., 2000; Sleeper et al., 2000) and along unmyelinated fibers and nerve endings in the cornea (Black and Waxman, 2002) and in the skin (Dib-Hajj et al., 2002; Rush et al., 2005). Nav1.9 is rare in Ab neurons (Dib-Hajj et al., 1998b; Tate et al., 1998b; Fjell et al., 1999a; Amaya et al., 2000; Fjell et al., 2000) but, when present, is restricted to nociceptive neurons (Fang et al., 2002). The presence of Nav1.9 at nerve endings is consistent with its predicted role of amplifying subthreshold stimuli (Herzog et al., 2001).

3.3 ROLE OF SODIUM CHANNELS IN TRAUMA-INDUCED PAINFUL NEUROPATHIES There is substantial evidence for a critical role of sodium channels in acquired and inherited painful neuropathies. A better understanding of the role of individual channels in chronic pain syndromes has been obtained from studies of animal models and human patients. The following sections will discuss our current knowledge about dysregulation and modulation of sodium channels in painful neuropathies. 3.3.1

Nerve Transection

Three of the most commonly used animal model for studying dysregulation of voltagegated sodium channels following nerve injury are sciatic nerve transection that entails cutting and tying off axons within the sciatic nerve at the midthigh level, spinal nerve ligation (SNL) that is caused by tight ligature of the spinal nerve originating from an individual DRG (Kim and Chung, 1992), and spared nerve injury (SNI) where two of the branches of the sciatic nerve, the tibial and the common peroneal, are cut sparing

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the third branch (sural) (Decosterd and Woolf, 2000). The main advantages of these models are that the injury is highly reproducible, permits easier application of exogenous factors, for example, neurotrophic factors or transformed cells that secrete desired factors, to the proximal nerve stump, and is amenable to assessment of behavioral pain responses. The application of exogenous factors has allowed investigations of the contribution of specific growth factors to the regulation of expression of voltage-gated sodium channels within DRG neurons (Dib-Hajj et al., 1998a; Boucher et al., 2000; Leffler et al., 2002). These injury models are similar in that nerve regeneration is prevented so that the severed axons cannot regrow to reach target tissues, either by threading the proximal end of the nerve into a silicon cuff (Waxman et al., 1994), or by effecting transection through a tight and permanent ligature of the spinal nerve (Kim et al., 2001, 2002) or the branches of the sciatic nerve (Decosterd and Woolf, 2000). The absence of axonal regeneration to peripheral tissues may not provide an exact parallel to clinical cases of trauma to human patients. However, each of these injury models results in the intermingling of injured and intact axons, which may lead to injury-mediated effects on the spared axons as will be discussed later. Neuropathic pain caused by transection of axons both in animal models and in human patients has a structural component at the site of injury and molecular and cellular components as well, which originate within the cell bodies of DRG neurons. The two components of the injury response are not mutually independent and reflect the interplay of neurons and various cell types, for example, immune cells, glia, and peripheral tissue cells such as keratinocytes. These cellular components constitute an integrated information unit that produces and transmits injury signals from the periphery to higher order centers within the CNS. For simplicity, we will separately discuss cellular, molecular, and structural components of response to injury. The physical transection of axons prevents the flow of electrical and chemical information between the neuronal cell bodies and their peripheral targets. At the molecular level, the physical transection of axons activates an injury response gene expression program that leads to an altered profile of ion channels, among other changes within DRG neurons (Costigan et al., 2002; Xiao et al., 2002), and secondorder neurons in the dorsal horn of the spinal cord (Yang et al., 2004a). The changes in gene expression underlie a new postinjury state of hyperexcitability of injured primary afferents and higher order neurons, which is manifested in neuropathic pain symptoms, for example, tactile allodynia and phantom pain (pain from an amputated limb). Although a multitude of gene products are dysregulated following trauma, studies from animal models and empirical data from human patients point to a key role of sodium channels in producing this neuronal hyperexcitability and the neuropathic pain state. Dynamic regulation of sodium channel expression after injury with some channels “turned off ” and others “turned on” is now a well-established contributor to hyperexcitability of DRG neurons (Fig. 3.3) (Waxman, 1999, 2001). Transcripts and protein levels of the Nav1.3 channel, which are detected only at exceedingly low levels in adult rat neurons, are upregulated following axotomy (Waxman et al., 1994; Dib-Hajj et al., 1996; Black et al., 1999). In contrast, transcripts and protein levels of Nav1.8 and Nav1.9 are significantly reduced in injured neurons (Dib-Hajj et al., 1996; Dib-Hajj

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FIGURE 3.3 Alterations in expression of Nav1.3, Nav1.8, and Nav1.9 in DRG neurons following peripheral transection of sciatic nerve. (A) RT-PCR analyses of control (C) and peripherally axotomized (A) DRG demonstrates upregulation of Nav1.3 and downregulation of Nav1.8 and Nav1.9 at 7–12 days following axotomy. (B) Contralateral (contra) and ipsilateral (ipsi) DRG reacted with isoform-specific antibodies for Nav1.3, Nav1.8, and Nav1.9 display an upregulation of Nav1.3 signal and a downregulation of immunofluorescent signal for Nav1.8 and Nav1.9 within DRG neurons. Scale bar: 50 mm. (C) Whole-cell patch-clamp recordings of control and axotomized small DRG neurons. (a) The time course for recovery from inactivation at 80 mV is faster in axotomized (open circles) than in control (filled circles) neurons. (b) Time constants for recovery from activation plotted as function of voltage. Time constants for axotomized (open circles) are smaller than that for control (filled circles) neurons. (c) and (d) Slowly inactivating (Nav1.8) and persistent (Nav1.9) TTX-resistant currents are reduced in small DRG neurons following peripheral axotomy. Scale bar: 100 mm. (Modified and reproduced with permission from Black et al. (1999), Dib-Hajj et al. (1996), Dib-Hajj et al. (1998a), and Sleeper et al. (2000).)

et al., 1998b; Sleeper et al., 2000; Decosterd et al., 2002), and those for Nav1.1, Nav1.6 and Nav1.7, the other voltage-gated sodium channels that are expressed in DRG neurons (Black et al., 1996), are reduced but to a lesser extent (Kim et al., 2001, 2002). Dysregulation of Nav1.3 following axotomy is not a generalized response to injury or a recapitulation of an embryonic expression profile. For example, transection of the

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sciatic nerve injures the peripheral axons of both sensory neurons of DRGs and primary motor neurons within the spinal cord; however, Nav1.3 is upregulated only in DRG neurons (Waxman et al., 1994; Dib-Hajj et al., 1996; Black et al., 1999) and not in motor neurons (Black and Waxman, unpublished observations). In addition, axotomy does not upregulate Nav1.2, another channel that is expressed within DRG neurons during embryogenesis (Waxman et al., 1994). Finally, transection of the central projections of DRG neurons (dorsal rhizotomy) does not cause an upregulation of Nav1.3 (Sleeper et al., 2000). The reduction of transcripts and protein for the Nav1.8 and Nav1.9 channels in axotomized DRG neurons is paralleled by the loss of the total TTX-R current in these neurons (Rizzo et al., 1995; Cummins and Waxman, 1997). The identification of the specific current profiles of Nav1.8 (Akopian et al., 1996; Sangameswaran et al., 1996; Akopian et al., 1999) and Nav1.9 (Cummins et al., 1999) permitted the unequivocal demonstration that both these channels and the sodium currents that they produce are downregulated after sciatic nerve transection (Cummins et al., 2000) and SNL (Gold et al., 2003) but not rhizotomy (Sleeper et al., 2000). The loss of Nav1.8 and Nav1.9 within injured DRG neurons has been confirmed in human patients suffering from traumatic injury to their ganglia (Coward et al., 2000). While the loss of these channels is likely to influence the electrogenic properties of injured DRG neurons, it is not clear whether their loss might contribute to hyperexcitability or increased spontaneous firing of these neurons (however, see discussion below on innervation of tissues by regenerating nerves). In contrast to the loss of the TTX-R currents in axotomized DRG neurons, the major effect on the TTX-S currents is a change in the repriming properties but with little effect on the current density. Axotomy of DRG neurons causes the loss of slowly repriming and the emergence of rapidly repriming TTX-S sodium current (Cummins and Waxman, 1997). Nav1.7 channels produce a slow-repriming TTX-S channels when expressed in native DRG neurons (Herzog et al., 2003) or in a heterologous expression system (Cummins et al., 1998). Thus, the loss of the slow-repriming TTX-S current in injured neurons (Cummins and Waxman, 1997) has been attributed to the reduction in Nav1.7 transcripts in DRG neurons from rats with SNL injury (Kim et al., 2002). The data from this animal study are consistent with studies on human patients with traumatic injury to their peripheral nerves, which shows that Nav1.7 immunostaining is significantly reduced in injured DRG neurons compared to control (Coward et al., 2001). The contribution of Nav1.6, the other TTX-S sodium channel that is known to be present in small DRG neurons (Black et al., 1996, 2002), to the sodium current profile in control and injured DRG neurons is not clear. Nav1.6 has been shown to rapidly reprime when expressed in DRG neurons, similar to its behavior in a heterologous expression system (Herzog et al., 2003). SNL injury reduces Nav1.6 transcript levels in rat DRG neurons (Kim et al., 2002), but the absence of a specific blocker or an intrinsic biophysical property that is not cell dependent hinders the determination of the effect of this injury on the Nav1.6 current. Thus, the contribution of Nav1.6 to DRG neuronal hyperexcitability after nerve injury remains less well understood than that of other channels.

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The emergence of a rapidly repriming TTX-S current in DRG neurons has been attributed, at least partially, to the upregulation of Nav1.3 following axotomy. Nav1.3 is virtually undetectable in normal adult DRG but is significantly upregulated following axotomy (Waxman et al., 1994; Black et al., 1996, 1999; Dib-Hajj et al., 1996) and in adult DRG neurons that have been dissociated and kept in culture for several days (Black et al., 1997). More important, Nav1.3 produces a rapidly repriming TTX-S current in DRG neurons and in a heterologous expression system (Cummins et al., 2001) with similar properties to those observed within injured DRG neurons (Cummins and Waxman, 1997). Injury-induced upregulation of Nav1.3 with its rapidly repriming properties and the ability to produce a significant current in response to the application of small, slow depolarizations (ramp stimulus) (Cummins et al., 2001) could support a lower threshold for single action potential firing and sustained highfrequency firing, which are hallmarks of neuronal hyperexcitability in injured neurons. The regenerative response of peripheral axons can produce a neuroma, which is a collection of de- and dysmyelinated axon sprouts and connective tissue at the proximal end of the transected axons (Fried et al., 1991). Neuromas have been shown to be sites of ectopic impulse generation and spontaneous firing of DRG neurons, which can lead to sensitization (for recent reviews see Black, ; Amir et al., 2006; Devor, 2006). For patients, painful neuromas can be worse than motor deficits associated with the injury. Interestingly, neuromas may not be painful, and in these cases, the accumulation of ankyrin G, which is normally clustered at nodes of Ranvier and initial segments, is reduced compared to that of painful neuromas or neurofibroma (Kretschmer et al., 2002, 2004). Treatment generally involves surgeries to resect the neuroma and implant the nerve end in different tissues (Vernadakis et al., 2003; Lewin-Kowalik et al., 2006). Early studies demonstrated accumulations of sodium channels at the distal tips of transected axons (Devor et al., 1989; England et al., 1996a). Importantly, Nav1.3 was specifically shown to accumulate within axon tips in the neuroma in rats (Black et al., 1999) in addition to its upregulation within the cell bodies of peripherally axotomized DRG neurons. The intensity of Nav1.3 immunoreactivity is highest at the distal region of the transected nerve, with only background levels of immunofluorescence at distances greater than 500–1000 mm proximal to this region. The accumulation of Nav1.3 within the neuroma suggests a molecular basis for the generation of ectopic discharges, which are known to emanate from this region (Scadding, 1981; Burchiel, 1984a, 1984b; Matzner and Devor, 1994). Consistent with a significant role of TTX-S sodium channel(s) in this process, application of
20 nM TTX to the neuroma significantly attenuates ectopic firing (Liu et al., 1999, 2001). The accumulation of Nav1.3 at the neuroma is coupled with the coaccumulation of the cell adhesion molecule contactin/F3 (Shah et al., 2004) (Fig. 3.4). Contactin has been shown to associate with several sodium channels and increase their current density both in heterologous expression systems (Kazarinova-Noyes et al., 2001; Shah et al., 2004) and in native DRG neurons (Rush et al., 2005). Previously, it was shown that contactin levels at the plasma membrane of hypothalamic neurons are regulated in an activity-dependent manner (Pierre et al., 2001). Therefore, a positive feedback loop could exist, leading to an increased contactin-mediated Nav1.3 surface expression at

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FIGURE 3.4 Nav1.3 and contactin are upregulated and colocalized in sciatic nerve neuroma— Images of uninjured control (contralateral) and transected (ipsilateral) sciatic nerve show that contactin and Nav1.3 accumulate within axon-like profiles within the neuroma that forms at the ligation site (arrow), with less immunostaining in proximal regions of the transected nerve. Colored images in the original publication (Shah et al., 2004) show that contactin and Nav1.3 exhibit extensive colocalization within the neuroma (not shown). (Modified and reproduced with permission from Shah et al. (2004).)

the neuroma, which could exacerbate neuropathic pain following nerve injury (Shah et al., 2004). In a limited number of human cases, Nav1.7 and Nav1.8 have been reported to accumulate within neuromas of trunk nerves (Coward et al., 2000; Kretschmer et al., 2002). Recently, Nav1.7 accumulation has been reported in painful neuromas of the lingual nerve but not in nonpainful neuromas of the same nerve (Bird et al., 2007). The accumulation of Nav1.8 at neuromas in human patients is in stark contrast to the findings from animal studies where sciatic nerve was studied. Similarly, the accumulation of Nav1.7 at sciatic nerve neuromas in animal models of injury has not been reported in the literature. Consistent with the downregulation of Nav1.8 and Nav1.9 mRNA and protein in DRG neurons following peripheral axotomy, these channels do not accumulate within the neuroma at 9–14 days following sciatic nerve transection (Black, ). The absence of Nav1.8 and Nav1.9 from neuromas is consistent with the blocking of ectopic firing at the neuroma by the application of 20 nM TTX (Liu et al., 1999, 2001), a TTX concentration that is several orders of magnitude lower than the Kd of these two TTX-R channels (Akopian et al., 1996; Cummins et al., 1999). Studies on human patients share the common weakness of small numbers of samples and a paucity of control tissue, two deficiencies that stress the need for caution in interpreting their outcomes.

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Chronic Constriction Injury

A chronic constriction injury (CCI) of the sciatic nerve is produced by placing loose ligatures around the nerve at midthigh level, which results in the comingling of injured and intact fibers for a long distance within the nerve and produces behavioral signs of spontaneous pain and cutaneous hyperalgesia in rodents (Bennett and Xie, 1988). DRG neurons subjected to CCI develop abnormal spontaneous activity and adrenergic sensitivity in the intact ganglion in vivo (Kajander et al., 1992; Xie et al., 1995) and in vitro (Zhang et al., 1997) or in dissociated DRG neurons (Petersen et al., 1996; Study and Kral, 1996). A linkage between Wallerian degeneration and hyperalgesia in CCI was confirmed by the demonstration that the onset of, and recovery from, hyperalgesia is delayed, with parallel delays in recruitment of macrophages to the site of injury, cytokine production, and degeneration of nerve fibers in the C57Bl/wld mouse strain compared to C57BL/6 (Myers et al., 1996; Sommer and Schafers, 1998). Most myelinated and unmyelinated axons (60–80%) undergo Wallerian degeneration following CCI (Basbaum et al., 1991; Carlton et al., 1991), an observation that has been verified functionally by the reduction in the anterograde axonal transport of acetyl cholinesterase (Filliatreau et al., 1994). CCI appears to produce effects that are a combination of axotomy and inflammation of nerve fibers. CCI triggers changes in sodium channel expression in both primary (DRG) and higher order sensory neurons, which may contribute to DRG neuron hyperexcitablity (Dib-Hajj et al., 1999; Hains et al., 2004a). The pattern of sodium channel dysregulation in DRG neurons at 14 days following CCI (Fig. 3.5) was similar but less extensive compared to that following sciatic nerve transection (Dib-Hajj et al., 1999), consistent with the observations that most of the sciatic nerve fibers are transected following this injury (Basbaum et al., 1991; Carlton et al., 1991). A reduction in transcripts and currents attributed to the TTX-R sodium channels Nav1.8 and Nav1.9 has been observed (Dib-Hajj et al., 1999; Decosterd et al., 2002), and an increase in the level of the TTX-S Nav1.3 channels is thought to contribute to the shift in the repriming properties of the TTX-S current in these neurons (Dib-Hajj et al., 1999). A similar reduction in the TTX-R current but with a hyperpolarizing shift in the voltagedependence of activation was independently reported (Kral et al., 1999). The difference might be attributable to recording of the currents in acutely dissociated DRG neurons from injured animals (Kral et al., 1999), compared to neurons in culture for 24 h (Dib-Hajj et al., 1999). At this point, however, it remains to be determined if contactin and Nav1.3 accumulate in injured axons proximal to the loose ligatures in the CCI model, similar to their accumulation at neuromas (Shah et al., 2004). The effect of CCI on the dysregulation of Nav1.8 and the contribution of this channel to DRG neuronal hyperexcitability, however, are a subject of controversy. Another study using CCI as an injury model reported no change in Nav1.8 transcripts and slowly inactivating TTX-R current density but a reduction in NaV1.8-specific immunolabeling within cell bodies and an accumulation of Nav1.8 in the sciatic nerve (Novakovic et al., 1998). An explanation that might reconcile the apparent discrepancy between this study and those discussed above would be to invoke an upregulation of Nav1.8 in intact nerves that comingle with degenerating axons of the sciatic nerve leading to an increased Nav1.8 immunostaining in these fibers. None of the reports on CCI measured levels of Nav1.8 in identified, uninjured DRG neurons following injury.

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FIGURE 3.5 Chronic constriction injury (CCI) of sciatic nerve upregulates Nav1.3 and downregulates Nav1.8 and Nav1.9 in DRG neurons. (a) RT-PCR products from control (C) and CCI DRG 14 days postsurgery exhibit a significant downregulation of Nav1.8 and Nav1.9 mRNA (Nav1.8: 512 bp; Nav1.9: 392 bp). Nav1.3 transcripts are upregulated in CCI neurons (Nav1.3: 412 bp). (b) In situ hybridization of small neurons from control and CCI DRG shows that hybridization signals for Nav1.8 and Nav1.9 are significantly reduced in CCI neurons, while hybridization signal for Nav1.3 is upregulated. Scale bar: 20 mm. (c) Recovery from inactivation is accelerated after CCI; repriming kinetics of TTX-sensitive current is significantly increased in CCI neurons compared to control neurons. Patch-clamp recording demonstrates a significant reduction in TTX-resistant currents in CCI neurons compared to control neurons. (Modified and reproduced with permission from Dib-Hajj et al. (1999).)

An upregulation of Nav1.8 in spared DRG neurons is in agreement with the finding that levels of substance P are significantly upregulated following CCI in DRG neurons with spared axons, possibly due to the exposure to inflammatory cytokines, for example, NGF, during Wallerian degeneration of sciatic nerve fibers (Ma and Bisby, 1998). NGF is known to increase levels of Nav1.8 within DRG neurons in vitro and in vivo (Dib-Hajj et al., 1998; Leffler et al., 2002). An elevation of Nav1.8 in spared neurons, whose axons comingle with damaged neighbors, would lead to neuronal hyperexcitability and might explain the lower incidence of ectopic firing following saphenous nerve transection in Nav1.8-null neurons (Roza et al., 2003) and the amelioration of neuropathic pain behavior in CCI animal injury models by knockdown of Nav1.8 using antisense oligonucleotides (Lai et al., 2002; Gold et al., 2003; Joshi et al., 2006). 3.3.3

Radicular Injury

Compression injuries either to the nerve root or to the DRG itself cause hyperexcitability that is manifested by periods of prolonged rapid firing and radicular pain (Howe et al., 1977). Radicular pain is a common clinical diagnosis that could have both mechanical and inflammatory components underlying its pathogenesis (Van Zundert et al., 2006). However, the molecular pathophysiology leading to neuronal

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hyperexcitability in compressed DRG neurons is not as well understood as that following SNL or axotomy. An animal model has been developed to investigate the molecular basis for DRG neuron hyperexcitability following chronic compression injury of DRG (CCD) (Hu and Xing, 1998). Compression of DRG neurons by placement of a metal rod in the intervertebral foramen, producing foramen stenosis and chronic compression of the ganglion, causes hyperexcitability of dissociated neurons or those in intact ganglion (Hu and Xing, 1998; Zhang et al., 1999; Ma and LaMotte, 2005). Spontaneous activity of DRG neurons following CCD is regulated by PKA (Hu et al., 2001), and these neurons demonstrate an enhanced response to inflammatory mediators (Song et al., 2003; Sun et al., 2006), both features that are consistent with neuronal hyperexcitability. CCD leads to dysregulation of sodium channel Nav1.9, albeit with less marked changes than SNL or axotomy (Abe et al., 2002). This injury, however, does not cause an increase in the levels of Nav1.3 transcripts or a significant reduction in the Nav1.8 channel transcript (Abe et al., 2002) unlike the effects of SNL and axotomy (see discussion above). However, recordings from identified cutaneous afferents revealed an 8 mV hyperpolarizing shift in voltage dependence of activation of TTX-S channels and an increase in the peak amplitude of the slowly inactivating TTX-R current, and a reduction in voltage-gated potassium current (Tan et al., 2006). The lack of correlation between electrophysiological changes and changes in channel transcript levels (Abe et al., 2002) suggests a posttranscriptional or posttranslational modulation of channels in the CCD injury model. The totality of the evidence strongly suggests that plasticity of expression, and possibly modulation, of sodium channels in compressed DRG neuron contribute to neuronal hyperexcitability in radicular pain.

3.4 ROLE OF SODIUM CHANNELS IN INFLAMMATION-INDUCED PAINFUL NEUROPATHIES Inflammation-induced pain behavior shows the same hallmark responses as those of neuropathic pain, including spontaneous activity of nociceptors, lowered threshold for action potential generation, and exaggerated evoked response to stimuli. The inflammation-induced pain symptoms are attenuated by treatment with sodium channel blockers (for a recent review see Amir et al., 2006), indicating a key role of these channels in neuronal hyperexcitability following tissue or nerve inflammation. The empirical data from human patients have been supported by animal studies, which demonstrate plasticity in the expression of sodium channels within nociceptors following inflammation of skin and muscle (Okuse et al., 1997; Tanaka et al., 1998; Tate et al., 1998; Gould et al., 1999; Black et al., 2004; Nassar et al., 2004) or viscera (Yoshimura et al., 2001; Bielefeldt et al., 2002; Beyak et al., 2004; Malykhina et al., 2004). Studies using knockout or knockdown models of specific sodium channels have identified Nav1.7, Nav1.8, and Nav1.9 as significant contributors to inflammation-induced pain. Inflammation of hindpaw in the rat causes an upregulation of Nav1.7 in DRG neurons that project to the inflamed area (Fig. 3.6) (Black et al., 2004; Gould

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FIGURE 3.6 Sodium channel protein expression and sodium currents in DRG neurons from saline-injected naive and carrageenan-injected rats. (a) Sections from saline and carrageenan-injected DRG were reacted with isoform-specific antibodies to Nav1.7 and Nav1.8. Sodium channels Nav1.7 and Nav1.8 exhibit greater immunolabeling in carrageenan neurons compared to saline-injected neurons. Microdensitometry quantitative analysis demonstrates that the immunofluorescent signals for Nav1.7 and Nav1.8 are significantly (*p 5 0.05) upregulated in small (5 25 mm) DRG neurons from carrageenan-injected DRG compared to saline-injected contralateral to carrageenan-injected (contra carrageenan) and saline-injected naive rat (saline) DRG. Scale bar: 50 mm. (b) Representative families of voltage-activated TTX-S (upper) and TTX-R (lower) sodium current traces for small neurons cultured from saline (left) and carrageenan (right) DRG are shown. Both TTX-S and TTX-R sodium current densities are significantly greater in carrageenan- than saline-injected small DRG neurons. (Modified and reproduced with permission from Tanaka et al. (1998) and Black et al. (2004).)

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et al., 2004). Nav1.7 shows the most robust increase in transcript and protein levels following carrageenan-induced inflammation of the hindpaw in rats, and this is accompanied by a significant increase in the amplitude of TTX-S current in DRG neurons (Black et al., 2004). Inflammation-induced increase in the levels of Nav1.7 transcript and protein might be regulated by the increased levels of NGF, which is known to upregulate Nav1.7 expression in cell lines (Toledo-Aral et al., 1995, 1997) and in DRG neurons (Gould et al., 2000). In the absence of specific channel blockers, it is difficult to definitively attribute the increase in the TTX-S current solely to increased Nav1.7 protein levels. However, knockdown of Nav1.7 in primary afferents has been shown to prevent inflammation-induced thermal hyperalgesia in mice injected with complete Freund’s adjuvant in their hindpaw (Yeomans et al., 2005). A direct role of Nav1.7 in inflammatory pain has been confirmed in studies using Nav1.7 knockout mice. Mice that lack Nav1.7 in DRG neurons show almost complete absence of inflammation-induced pain (Nassar et al., 2004). Additional evidence has come from patients with the painful neuropathy inherited erythromelalgia (which will be discussed in a separate section) who show swelling, erythema, and local increase in skin temperature (Novella et al., 2007), symptoms that are common to inflammatory conditions, as a result of gain-of-function mutations of Nav1.7. Thus, Nav1.7 appears to play an important role in inflammation-induced pain. Chronic inflammation causes an increase in the level of Nav1.8 channels in affected DRG neurons, which may contribute to neuronal excitability leading to pain. Carrageenan-induced inflammation of rat hindpaw causes an increase in the Nav1.8 current 4 days postinjection (Tanaka et al., 1998), which is paralleled by an increase in the intensity of immunoreactivity of DRG neurons to Nav1.8-specific antibody (Black et al., 2004). CFA injection into rat hindpaw does not increase levels of Nav1.8 transcripts or protein within DRG neurons after 48 h (Okuse et al., 1997) but appears to induce an increase in the translocation of the channel protein to a larger number of myelinated and unmyelinated axons in sciatic nerve (Coggeshall et al., 2004). Colitis, which is induced 10 days after injection of trinitro genzone sulfonic acid causes an increase in the Nav1.8 current density 10 days postinjection (Beyak et al., 2004). Reducing the levels of the channel in DRG neurons by the administration of anti-Nav1.8 antisense oligonucleotides ameliorates inflammatory pain induced by PGE2 (Khasar et al., 1998) or CFA (Porreca et al., 1999; Joshi et al., 2006). Similarly, acetic acid-induced urinary bladder pain is attenuated using anti-Nav1.8 antisense oligonucleotide treatment (Yoshimura et al., 2001). Thus, an increase in Nav1.8 levels in DRG neurons can contribute to the development of inflammatory pain. Chronic or acute application of inflammatory mediators, injected subcutaneously in vivo or applied directly to DRG neurons in culture, causes an increase in the amplitude of the TTX-R slowly inactivating Nav1.8 current. NGF, which functions as an inflammatory cytokine in addition to being a trophic factor (Hefti et al., 2006), contributes to the chronic upregulation of Nav1.8 levels within DRG neurons under inflammatory conditions. NGF treatment triggers an increase in Nav1.8 expression in DRG neurons both in vivo (Dib-Hajj et al., 1998a; Fjell et al., 1999b; Leffler et al., 2002) and in vitro (Fjell et al., 1999b; Cummins et al., 2000). Acute application

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of inflammatory mediators PGE2 (England et al., 1996; Gold et al., 1996), NGF and its second messenger ceramide (Zhang et al., 2002), and endothelin-1 (Zhou et al., 2002) have been shown to increase the TTX-R Nav1.8 current density, and the increase is accompanied by a hyperpolarizing shift in voltage dependence of activation of the channel. However, activation of p38 MAP kinase in DRG neurons, either by treatment with TNF-a (Jin and Gereau, 2006), a major inflammatory cytokine, or by anisomycin (Hudmon, 2008) to induce a stress response, increases the amplitude of the TTX-R current but without a change in the gating properties of the channel. The increase in Nav1.8 current density by activated p38 MAPK is mediated by the direct phosphorylation of two residues in loop 1 of Nav1.8 (Hudmon, 2008). Inflammation of visceral organs has been associated with an increase in the current density of the slowly inactivating TTX-R current but without a hyperpolarizing shift in the voltage dependence of activation (Yoshimura et al., 2001; Bielefeldt et al., 2002; Malykhina et al., 2004). These observations suggest that diverse inflammatory modalities may regulate the Nav1.8 current in different ways. Alternatively, altered gating properties of Nav1.8 in some studies may reflect the use of different recording conditions. Studies using Nav1.8-null mice confirm the important role of this channel in painful neuropathies. Mice that lack Nav1.8 demonstrate a delayed development of carrageenan-evoked (Akopian et al., 1999), an attenuated NGF-evoked (Kerr et al., 2001), thermal hyperalgesia, and reduced visceral pain responses to inflammatory mediators and referred hyperalgesia (Laird et al., 2002). Nav1.8-null mice also show deficits in visceral pain following parasite-induced chronic inflammation (Hillsley et al., 2006), which is consistent with the presence of Nav1.8 in sensory DRG neurons innervating the colon (Gold et al., 2002). These findings are best explained by the role of this channel in rendering neurons hyperexcitable (Rush et al., 2006). The contribution of Nav1.9 to inflammatory pain, while not as well understood as that of Nav1.7 and Nav1.8, has begun to be clarified by studies of mice that lack this channel, in which the persistent Nav1.9 TTX-R current is absent. An increase in the level of Nav1.9 transcripts was reported 7 days after CFA injection into the hindpaw of rats (Tate et al., 1998). PGE2, which acts on this channel via a G-protein-coupled mechanism, increases the current density of Nav1.9 (Rush and Waxman, 2004). The increase in the current density of Nav1.9 via G-proteins has been linked to a reduction in the threshold for action potential firing and an increase in spontaneous firing (Baker et al., 2003). This effect of Nav1.9 on neuronal excitability is consistent with predictions from computer simulations that this channel contributes to setting resting membrane potential and amplifying subthreshold stimuli (Herzog et al., 2001). In agreement with a role of Nav1.9 in inflammatory pain, heterozygotes and Nav1.9-null mice show deficits in the late phase of formalin-induced pain, and Nav1.9null mice show a reduction in the duration of thermal hyperalgesia following carrageenan or CFA injection into the hindpaw (Priest et al., 2005). Recently, similar findings in another Nav1.9-null mouse were reported in another study, which extended the analysis to involve several inflammatory mediators that act through distinct second

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messenger systems (Amaya et al., 2006). However, there was no difference in pain behavior between wild-type and Nav1.9-null mice when PGE2 was injected into the subarachnoid space of spinal cord, suggesting that Nav1.9 does not contribute to direct central sensitization (Amaya et al., 2006), consistent with the restriction of the channel’s expression to peripheral sensory neurons (Dib-Hajj et al., 1998b; Tate et al., 1998). Furthermore, NGF-mediated effects were not different between wild-type and Nav1.9-null mice (Amaya et al., 2006), consistent with the expression of Nav1.9 in DRG neurons that respond to glial-derived neurotrophic factor but not NGF (Cummins et al., 2000; Fjell et al., 1999a). However, unlike Nav1.8-null mice, yet another Nav1.9-null mouse model showed no deficits in nematode-induced inflammatory visceral pain (Hillsley et al., 2006). It is noteworthy that Nav1.9 has been detected in myenteric sensory neurons (Rugiero et al., 2003) but not in afferents innervating the bladder (Black et al., 2003). Taken together, these data support an important role of Nav1.9 in some (but not all) inflammatory pain conditions.

3.5 ROLE OF SODIUM CHANNELS IN PAINFUL DIABETIC NEUROPATHY Sodium channel dysregulation has been investigated in diabetic neuropathy, which was evoked in rats by treatment with streptozotocin (STZ) that resulted in tactile allodynia 6 weeks following the injection of the drug (Craner et al., 2002). Dysregulated expression of sodium channels (Fig. 3.7) was evident in diabetic rats 8 weeks after the onset of allodynia with significant increases in transcript levels in small DRG neurons (525 mm) of Nav1.3 and Nav1.6, no change in Nav1.1, Nav1.7 and Nav1.9, and a reduction in Nav1.8 (Craner et al., 2002). Protein levels generally mirrored transcript levels in the diabetic neurons, except for a small increase in Nav1.9 protein (Craner et al., 2002). Notably, the expression of Nav1.9 was shown to increase substantially in large DRG neurons. Nav1.7 was reported to increase as judged by immunoblot assay in another study (Hong et al., 2004). Dysregulation of sodium channel transcript and protein levels in diabetic neurons is reflected in changes to the TTX-S and TTX-R sodium currents in these neurons (Hong et al., 2004). Electrophysiological studies showed an increase in the TTX-S sodium current, consistent with the upregulation of Nav1.3, Nav1.6, and Nav1.7 channels (as discussed above). Importantly, there was a significant increase in the current in response to a small, slow depolarization (ramp stimulus) of the diabetic neuron (Hong et al., 2004). The increase in ramp current is consistent with an upregulation of Nav1.3 and/or Nav1.7 channels since both the channels have been shown to produce a robust ramp current both in DRG neurons and in heterologous expression systems (Cummins et al., 1998; Cummins et al., 2001; Herzog et al., 2003). The increase in ramp currents in diabetic neurons may contribute to a lowered threshold for firing in these neurons. The electrophysiological studies, however, have also shown that STZ-induced diabetes increases the slowly inactivating TTX-R current and shifts the voltage

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FIGURE 3.7 Sodium channel expression in control, 1-week, and 8-week allodynic DRG neurons in the rat streptozotocin model of diabetes. Nav1.3 and Nav1.6 immunoreactivity is increased in both 25 and 425 mm diameter diabetic DRG neurons at 1- and 8-week allodynia compared to control neurons. Nav1.8 immunolabeling in both neuron size classes at 1-week allodynia is similar to that exhibited by control neurons; in contrast, 425 mm DRG neurons display decreased Nav1.8 immunoreactivity at 8-week allodynia. Scale bar: 25 mm. (Modified and reproduced with permission from Craner et al. (2002).)

dependence of activation and steady-state inactivation in a hyperpolarizing direction (Hong et al., 2004), in disagreement with the reduction in transcript and protein levels of this channel (Craner et al., 2002; Hong et al., 2004). In an attempt to reconcile this paradoxical finding, Hong et al. (2004) show an elevated level of serine/threonine phosphorylation of Nav1.8 in diabetic neurons. Phosphorylation of Nav1.8 at serine residues in L1 causes an increase in the peak current of this channel in a PKA- and PKC-dependent manner (Fitzgerald et al., 1999; Vijayaragavan et al., 2004) and by stress activated mitogen-activated protein kinase (MAPK) p38 (Hudmon, 2008). Thus, the reduction of the total protein of Nav1.8 in DRG neurons appears to be overcompensated by the modulation of the channel by phosphorylation, which may enhance the stability of the channel at the cell surface and/or alter its gating to increase single-channel properties. The increased levels of Nav1.8 current are predicted to enhance neuronal excitability. The functional consequences of diabetic-induced changes to the persistent TTX-R Nav1.9 current (Cummins et al., 1999), however, remain to be examined.

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FIGURE 3.8 Nav1.3 expression and pain-related phenomena after spinal cord injury (SCI). In the lumbar dorsal horn of intact animals (a), Nav1.3 protein expression is very low. In contrast, 28 days following T9 SCI, Nav1.3 expression is upregulated in lumbar dorsal horn. At

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3.6 SODIUM CHANNEL DYSREGULATION AND CENTRAL SENSITIZATION AND CENTRAL PAIN Peripheral injury leads to sensory neuronal hyperexcitability that propagates postsynaptically to second- and third-order nociceptive neurones in the CNS; this process has been termed as central sensitization, and it shares many aspects of LTP (Ji, 2004; Woolf, 2004; Campbell and Meyer, 2006). The release of neurotransmitters from primary afferent terminals in the superficial lamina of the dorsal horn is traditionally thought to be a major determinant of central sensitization (Hunt and Mantyh, 2001). The LTP model of inflammatory hyperalgesia is supported by recent findings showing that low-level activity of presynaptic nociceptive fibers, similar to the activity during inflammation of peripheral targets, can lead to central sensitization, suggesting the presence of a synaptic amplifier in the dorsal horn (Ikeda et al., 2006). In this chapter, we will concentrate on the role of sodium channel dysregulation, which also appears to contribute to central sensitization and central pain. Sodium channel dysregulation in second- and third-order nociceptive neurons has been shown to contribute to central sensitization following peripheral nerve injury (Fig. 3.8). Specifically, CCI of sciatic nerve causes the upregulation of sodium channel Nav1.3 in primary sensory neurons, and this expression plasticity is extended postsynaptically into higher order neurons (Hains et al., 2003). In additional, contusion spinal cord injury (SCI) results in the upregulation of Nav1.3 in neurons along the spinothalamic tract, which are located caudal to the site of injury (Hains et al., 2003). The available evidence suggests that expression of Nav1.3 may contribute to hyperexcitability of these neurons, enlargement of peripheral receptive fields, and central pain (Waxman and Hains, 2006). The dysregulated expression of Nav1.3 in spinal cord neurons following peripheral injury or spinal cord injury suggests that sodium channel blockers may be effective therapeutic agents after SCI. Indeed, phenytoin treatment provides neuroprotection and improves functional outcome in an experimental model of 3

this timepoint, antisense (AS) oligodeoxynucleotides generated against Nav1.3 or its mismatch (MM) sequence were topically administered to the lumbar spinal cord. Extracellular unit recordings of multireceptive neurons revealed hyperresponsiveness to application of brush (BR), press (PR), pinch (PI), increasing strength von Frey filaments (0.39 g, 1.01 g, 20.8 g), and a noxious thermal stimulus (47  C) in MM-treated animals after SCI (c). In AS-receiving animals, evoked hyperresponsiveness of dorsal horn neurons after SCI was reduced. Nav1.3 expression was similarly low in the ventral posterolateral (VPL) nucleus of the thalamus in intact animals (e). In contrast, 4 weeks after SCI, Nav1.3 expression was upregulated in the VPL (f), but not adjacent structures. Unit recordings of VPL neurons following SCI and MM administration revealed no changes in hyperresponsiveness following stimulation of hindpaw receptive fields (g), however, AS administration resulted in reduced evoked responsiveness of VPL neurons (h). Lumbar intrathecal administration of AS, but not MM, resulted in a partial restoration of paw withdrawal thresholds to mechanical stimulation of the hindpaw (i), as well as paw withdrawal latencies to noxious thermal stimulation (j). Dotted line represents mechanical and thermal thresholds in intact animals. *p 5 0.05 SCI versus SCI þ AS. (Modified and reproduced with permission from Hains et al. (2004a, 2005).)

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contusion SCI (Hains et al., 2004). In addition, several studies have shown that the knockdown of Nav1.3 in central nocieptive neurons following peripheral (CCI injury model) or contusion SCI reduces hyperexcitability of these neurons and attenuates neuropathic pain behavior (Hains et al., 2003, 2004a, 2006). Two articles, however, have presented contradicting findings to the prevailing view of a key role of Nav1.3 in neuropathic pain (Lindia et al., 2005; Nassar et al., 2006). Mice with global or DRG-specific knockout of Nav1.3 develop normally, are fertile, and, surprisingly, have been reported to demonstrate normal neuropathic pain behavior after nerve injury (Nassar et al., 2006). It is possible though that either global or DRG-specific deletion of Nav1.3 leads to unexpected gene expression adjustments that might influence the behavior of the mice. This view is consistent with the modest effect of knocking out Nav1.8 on neuropathic pain behavior (Akopian et al., 1999) despite the fact that Nav1.8 contributes most of the current underlying the depolarizing phase of action potential in the neurons where it is expressed (Renganathan et al., 2001; Blair and Bean, 2002). Alternatively, the role of Nav1.3 in rendering primary and higher order neurones hyperexcitable might be redundant, and it may be replaced by another channel in the knockout mice. Lindia et al. (2005) have confirmed that SNL causes an upregulation of Nav1.3 within DRG neurons and have reported that the administration of specific antisense oligonucleotides intrathecally reduces the Nav1.3 immunoreactivity by 50% but does not ameliorate the mechanical or cold allodynia typical of this animal model (Lindia et al., 2005). In contrast, studies by Hains et al. (2003, 2004a, 2006) have reported near-complete reversal of injury-induced Nav1.3 upregulation in secondand third-order neurons following intrathecal administration of Nav1.3 antisense oligonucleotides. It is possible that a 50% reduction in Nav1.3 is not enough to attenuate neuropathic pain behavior. Alternatively, differences in antisense chemistry, synthesis, purification, dose, or delivery methods could be significant factors that affect the outcome of these studies. In addition, because none of these studies have investigated global gene expression following the antisense administration, it is possible that off-target effects, either pro- or antinociceptive, might have lead to the different outcomes. A direct test of the contribution of overexpression of Nav1.3 on excitability of DRG neurons could shed light on the potential role of this channel in painful neuropathies.

3.7 SODIUM CHANNELS AND INHERITED PAINFUL NEUROPATHIES Inherited painful peripheral neuropathies may have diverse etiologies. Mutations in sodium channel Nav1.7 have been identified in patients suffering from inherited erythromelalgia (IEM) (Waxman and Dib-Hajj, 2005; Dib-Hajj, 2006) and paroxysmal extreme pain disorder (PEPD) (Fertleman et al., 2006). Recently, individuals with congenital and complete loss of Nav1.7 have been reported to be “indifferent” to pain while showing no deficits in other sensory modalities (Cox et al., 2006). Thus, similar to acquired sodium channelopathies following trauma or inflammation, some inherited painful disorders are sodium channelopathies.

SODIUM CHANNELS AND INHERITED PAINFUL NEUROPATHIES

3.7.1

89

Inherited Erythromelalgia

Genetic studies in human patients with early-onset inherited erythromelalgia, a heritable disorder characterized by severe painful symptoms of the extremities that are accompanied by reddening of skin, have linked it to mutations in Nav1.7 (Waxman and Dib-Hajj, 2005; Dib-Hajj, 2006). Early-onset primary erythromelalgia (also known as erythermalgia) is a lifelong, autosomal dominant disorder that has been linked to missense mutations in Nav1.7 (Yang et al., 2004b; Dib-Hajj et al., 2005; Drenth et al., 2005; Michiels et al., 2005; Han et al., 2006; Harty et al., 2006). All these mutations have been shown to lower the threshold for channel activation, and most cause an increase in the amplitude of ramp currents and slowed deactivation kinetics (Cummins et al., 2004; Dib-Hajj et al., 2005; Choi et al., 2006; Han et al., 2006; Harty et al., 2006; Lampert et al., 2006). The introduction of these mutant channels into DRG neurons, such as Nav1.7/A863P, renders these neurons hyperexcitable by lowering the threshold for single action potential firing and increases the number of action potentials in response to a graded stimulus (Fig. 3.9) (Dib-Hajj et al., 2005; Harty et al., 2006; Rush et al., 2006). The expression of some mutant Nav1.7 channels in DRG neurons has been associated with a depolarization of the resting membrane potential of these neurons (Harty et al., 2006; Rush et al., 2006). This effect of mutant Nav1.7 channels brings the resting membrane potential of DRG neurons closer to the activation threshold voltage of Nav1.8 (Akopian et al., 1996; Sangameswaran et al., 1996). Depolarization of neuronal resting membrane potential in neurons that do not normally produce Nav1.8 channels is predicted to cause these neurons to become hypoexcitable because of resting inactivation of the TTX-S channels in these cells, which have hyperpolarized voltage dependence of activation and inactivation compared to Nav1.8 (Rush et al., 2007). Indeed, Nav1.7 mutant-mediated neuronal hyperexcitability appears to be dependent upon the presence of Nav1.8 because expression of L858H mutant Nav1.7 in superior cervical ganglion (SCG) neurons, which do not express endogenous Nav1.8, renders these neurons hypoexcitable (Rush et al., 2006). The coexpression of L858H Nav1.7 mutant and Nav1.8 in SCG neurons restores excitability of these neurons to near-normal levels although the L858H mutation depolarizes these cells (Rush et al., 2006), showing that the response to depolarization is conditioned by the absence (SCG neurons) or presence (DRG neurons) of Nav1.8. However, depolarization of the resting membrane potential does not explain all aspects of the behavior of DRG neurons housing Nav1.7 mutant channels (Harty et al., 2006), suggesting that other changes in the properties of mutant Nav1.7 channels may contribute to the hyperexcitability of DRG neurons in patients with inherited erythromelalgia. 3.7.2

Paroxysmal Extreme Pain Disorder

Recently, another inherited painful syndrome, paroxysmal extreme pain disorder (PEPD), previously known as familial rectal pain (Bednarek et al., 2005; Fertleman and Ferrie, 2006), has been linked to a different set of mutations in Nav1.7 (Fertleman et al., 2006). PEPD is manifested as severe pain and flushing in the lower body in infants during bowel movement, transforming with age to ocular and mandibular

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FIGURE 3.9 Nav1.7/A863P mutation decreases action potential threshold and increases frequency of firing in small, current-clamped DRG neurons. (a) Responses of a current-clamped DRG neuron transfected with WT Nav1.7 DNA to a series of subthreshold and suprathreshold depolarizing current steps. Starting at subthreshold stimulus intensity, the current amplitude was increased in 5 pA increments to intensity well beyond threshold. RMP for this cell was 55 mV and threshold was 310 pA. A DRG neuron expressing WT channels responds to a 1 s depolarizing current step, that is one, two, and three times the current threshold for action potential generation by the firing of up to two spikes (at 3 threshold). (b) The same threshold protocol applied to a DRG neuron transfected with the A863P mutant DNA elicits AP with a smaller current injection. RMP for this cell was 45 mVand threshold was 95 pA. Arrows with numbers indicate the current step amplitude used to elicit the labeled response. A DRG neuron expressing A863P mutant channels responds to a 1 s depolarizing current step, that is, one, two, and three times the current threshold for action potential generation by the firing of up to 11 spikes (at 3 threshold). (Modified and reproduced with permission from Harty et al. (2006).)

pain (Fertleman and Ferrie, 2006). Three mutations were characterized by whole-cell patch clamp analysis and were shown to impair fast inactivation of the mutant channels but with no effect on channel activation (Fertleman et al., 2006). Impaired fast inactivation allows more sodium current to flow through the mutant channel and is

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predicted to increase DRG neuron hyperexcitability. Interestingly, patients with PEPD tend to respond favorably to treatment with the anticonvulsant drug carbamazepine (Fertleman et al., 2006), unlike patients with inherited erythromelalgia who are refractory to long-term treatment with nonselective sodium channel blockers (Novella et al., 2007). 3.7.3

Congenital Insensitivity to Pain

Congenital insensitivity to experience pain is a very rare clinical phenotype. Recently, this trait has been shown to be inherited in an autosomal recessive manner in three consanguineous families from Pakistan, and nonsense mutations in Nav1.7 have been genetically linked to this phenotype (Cox et al., 2006). Patients missing functional Nav1.7 channels do not experience pain from inserting sharp objects in their hands or walking on hot surfaces (burning coal) or after bone fracture and tongue biting, but without deficits in other sensory modalities, for example, sensation of warmth or touch (Cox et al., 2006). Parents are asymptomatic, suggesting that SCN9A does not cause haploinsufficiency. The truncated channel protein produces no sodium current when expressed in the mammalian cell line HEK 293, suggesting a molecular pathophysiological cause for the phenotype (Cox et al., 2006). These data are in agreement with the results from animal studies using mice that lack Nav1.7 (Nassar et al., 2004), although the human and mouse cases differ in that the neuropathic component of pain in the animal model is intact (Nassar et al., 2005) whereas it appears to be missing in the patients (Cox et al., 2006).

3.8 CONCLUSIONS Sodium channelopathies underlie acquired and inherited neuropathies, some forms of epilepsy, and diseases of skeletal and cardiac muscles (Waxman, 2001; Lai et al., 2004; Viswanathan and Balser, 2004; Wood et al., 2004; George, 2005; Meisler and Kearney, 2005; Moss and Kass, 2005; Waxman and Dib-Hajj, 2005; Cannon, 2006; Venance et al., 2006; Waxman and Hains, 2006). Dysregulated expression of several sodium channels under a variety of pathological conditions induced by trauma or inflammation, and metabolic disorders, such as diabetes, has been linked to chronic pain, and blocking of sodium channels has been shown to ameliorate, or at least partially ameliorate, pain symptoms in some clinical studies of neuropathic pain in humans (Waxman, 2001; Lai et al., 2004; Wood et al., 2004; Waxman and Hains, 2006). Importantly, existing sodium channel blockers can be effective in treatment of some cases of neuropathic pain but suffer from undesirable side effects because they are not isoform selective (Rice and Hill, 2006). The contribution of individual channels to the pain state, however, is only now becoming better understood. As the roles of different channel isoforms under pathological conditions are elucidated, the identification of sodium channel isoforms that play dominant roles in pain should contribute to mechanism-based diagnosis and identify potential targets for treatment.

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REFERENCES Abe M, Kurihara T, Han W, Shinomiya K, Tanabe T, 2002. Changes in expression of voltagedependent ion channel subunits in dorsal root ganglia of rats with radicular injury and pain. Spine 27: 1517–1524 discussion 1525. Abriel H, Kass RS, 2005. Regulation of the voltage-gated cardiac sodium channel Nav1.5 by interacting proteins. Trends Cardiovasc Med 15: 35–40. Akopian AN, Sivilotti L, Wood JN, 1996. A tetrodotoxin-resistant voltage-gated sodium channel expressed by sensory neurons. Nature 379: 257–262. Akopian AN, Souslova V, England S, Okuse K, Ogata N, Ure J, Smith A, Kerr BJ, McMahon SB, Boyce S, Hill R, Stanfa LC, Dickenson AH, Wood JN, 1999. The tetrodotoxinresistant sodium channel SNS has a specialized function in pain pathways. Nat Neurosci 2: 541–548. Amaya F, Decosterd I, Samad TA, Plumpton C, Tate S, Mannion RJ, Costigan M, Woolf CJ, 2000. Diversity of expression of the sensory neuron-specific TTX-resistant voltage-gated sodium ion channels SNS and SNS2. Mol Cell Neurosci 15: 331–342. Amaya F, Wang H, Costigan M, Allchorne AJ, Hatcher JP, Egerton J, Stean T, Morisset V, Grose D, Gunthorpe MJ, Chessell IP, Tate S, Green PJ, Woolf CJ, 2006. The voltage-gated sodium channel Nav1.9 is an effector of peripheral inflammatory pain hypersensitivity. J Neurosci 26: 12852–12860. Amir R, Argoff CE, Bennett GJ, Cummins TR, Durieux ME, Gerner P, Gold MS, Porreca F, Strichartz GR, 2006. The role of sodium channels in chronic inflammatory and neuropathic pain. J Pain 7: S1–29. Arroyo EJ, Xu T, Grinspan J, Lambert S, Levinson SR, Brophy PJ, Peles E, Scherer SS, 2002. Genetic dysmyelination alters the molecular architecture of the nodal region. J Neurosci 22: 1726–1737. Baker MD, Chandra SY, Ding Y, Waxman SG, Wood JN, 2003. GTP-induced tetrodotoxinresistant Naþ current regulates excitability in mouse and rat small diameter sensory neurones. J Physiol Lond 548: 373–382. Basbaum AI, Gautron M, Jazat F, Mayes M, Guilbaud G, 1991. The spectrum of fiber loss in a model of neuropathic pain in the rat: an electron microscopic study. Pain 47: 359–367. Beckh S, Noda M, Lubbert H, Numa S, 1989. Differential regulation of three sodium channel messenger RNAs in the rat central nervous system during development. EMBO J 8: 3611– 3616. Bednarek N, Arbues AS, Motte J, Sabouraud P, Plouin P, Morville P, 2005. Familial rectal pain: a familial autonomic disorder as a cause of paroxysmal attacks in the newborn baby. Epileptic Disord 7: 360–362. Bennett GJ, Xie YK, 1988. A peripheral mononeuropathy in rat that produces disorders of pain sensation like those seen in man. Pain 33: 87–107. Beyak MJ, Ramji N, Krol KM, Kawaja MD, Vanner SJ, 2004. Two TTX-resistant Naþ currents in mouse colonic dorsal root ganglia neurons and their role in colitis-induced hyperexcitability. Am J Physiol Gastrointest Liver Physiol 287: G845–G855. Bielefeldt K, Ozaki N, Gebhart GF, 2002. Mild gastritis alters voltage-sensitive sodium currents in gastric sensory neurons in rats. Gastroenterology 122: 752–761. Bird EV, Robinson PP, Boissonade FM, 2007. Nav1.7 sodium channel expression in human lingual nerve neuromas. Arch Oral Biol 52: 494–502.

REFERENCES

93

Black JA, Cummins TR, Plumpton C, Chen YH, Hormuzdiar W, Clare JJ, Waxman SG, 1999. Upregulation of a silent sodium channel after peripheral, but not central, nerve injury in DRG neurons. J Neurophysiol 82: 2776–2785. Black JA, Cummins TR, Yoshimura N, de Groat WC, Waxman SG, 2003. Tetrodotoxinresistant sodium channels Nav1.8/SNS and Nav1.9/NaN in afferent neurons innervating urinary bladder in control and spinal cord injured rats. Brain Res 963: 132–138. Black JA, Dib-Hajj S, McNabola K, Jeste S, Rizzo MA, Kocsis JD, Waxman SG, 1996. Spinal sensory neurons express multiple sodium channel alpha-subunit mRNAs. Brain Res Mol Brain Res 43: 117–131. Black JA, Langworthy K, Hinson AW, Dib-Hajj SD, Waxman SG, 1997. NGF has opposing effects on Naþ channel III and SNS gene expression in spinal sensory neurons. Neuroreport 8: 2331–2335. Black JA, Liu S, Tanaka M, Cummins TR, Waxman SG, 2004. Changes in the expression of tetrodotoxin-sensitive sodium channels within dorsal root ganglia neurons in inflammatory pain. Pain 108: 237–247. Black JA, Renganathan M, Waxman SG, 2002. Sodium channel Nav1.6 is expressed along nonmyelinated axons and it contributes to conduction. Brain Res Mol Brain Res 105: 19–28. Black JA, Waxman SG, 2002. Molecular identities of two tetrodotoxin-resistant sodium channels in corneal axons. Exp Eye Res 75: 193–199. Black JA, Waxman SG, Smith KJ, 2006. Remyelination of dorsal column axons by endogenous Schwann cells restores the normal pattern of Nav1.6 and Kv1.2 at nodes of Ranvier. Brain 129: 1319–1329. Black J, Hains B, Dib-Hajj SD Waxman SG, 2005. Voltage-gated sodium channels and pain associated with nerve injury and neuropathies. In: Coward K, Baker MD (Eds), Sodium Channels, Pain, and Analgesia, Basel, Switzerland: Birkhauser Verlag, pp. 1–21. Blair NT, Bean BP, 2002. Roles of tetrodotoxin (TTX)-sensitive Naþ current, TTX-resistant Naþ current, and Ca2þ current in the action potentials of nociceptive sensory neurons. J Neurosci 22: 10277–10290. Blum R, Kafitz KW, Konnerth A, 2002. Neurotrophin-evoked depolarization requires the sodium channel Na(V)1.9. Nature 419: 687–693. Boiko T, Rasband MN, Levinson SR, Caldwell JH, Mandel G, Trimmer JS, Matthews G, 2001. Compact myelin dictates the differential targeting of two sodium channel isoforms in the same axon. Neuron 30: 91–104. Bongenhielm U, Nosrat CA, Nosrat I, Eriksson J, Fjell J, Fried K, 2000. Expression of sodium channel SNS/PN3 and ankyrin(G) mRNAs in the trigeminal ganglion after inferior alveolar nerve injury in the rat. Exp Neurol 164: 384–395. Boucher TJ, Okuse K, Bennett DL, Munson JB, Wood JN, McMahon SB, 2000. Potent analgesic effects of GDNF in neuropathic pain states. Science 290: 124–127. Burchiel KJ, 1984a. Effects of electrical and mechanical stimulation on two foci of spontaneous activitywhichdevelopinprimary afferentneuronsafterperipheralaxotomy.Pain 18:249–265. Burchiel KJ, 1984b. Spontaneous impulse generation in normal and denervated dorsal root ganglia: sensitivity to alpha-adrenergic stimulation and hypoxia. Exp Neurol 85: 257–272. Burgess DL, Kohrman DC, Galt J, Plummer NW, Jones JM, Spear B, Meisler MH, 1995. Mutation of a new sodium channel gene, Scn8a, in the mouse mutant ‘motor endplate disease’. Nat Genet 10: 461–465. Campbell JN, Meyer RA, 2006. Mechanisms of neuropathic pain. Neuron 52: 77–92.

94

VOLTAGE-GATED SODIUM CHANNELS

Cannon SC, 2006. Pathomechanisms in channelopathies of skeletal muscle and brain. Annu Rev Neurosci 29: 387–415. Cantrell AR, Catterall WA, 2001. Neuromodulation of Naþ channels: an unexpected form of cellular plasticity. Nat Rev Neurosci 2: 397– 407. Carlton SM, Dougherty PM, Pover CM, Coggeshall RE, 1991. Neuroma formation and numbers of axons in a rat model of experimental peripheral neuropathy. Neurosci Lett 131: 88–92. Catterall WA, 2000. From ionic currents to molecular mechanisms: the structure and function of voltage-gated sodium channels. Neuron 26: 13–25. Catterall WA, Goldin AL, Waxman SG, 2005. International Union of Pharmacology. XLVII. Nomenclature and structure-function relationships of voltage-gated sodium channels. Pharmacol Rev 57: 397–409. Choi JS, Dib-Hajj SD, Waxman SG, 2006. Inherited erythermalgia: limb pain from an S4 charge-neutral Na channelopathy. Neurology 67: 1563–1567. Chung JM, Dib-Hajj SD, Lawson SN, 2003. Sodium channel subtypes and neuropathic pain. In: Dostrovsky JO, Carr DB, Koltzenberg M (Eds), Proceedings of the 10th Congress in Pain Research and Management, Vol. 24. Seatle: IASP Press, pp 99–114. Coggeshall RE, Tate S, Carlton SM, 2004. Differential expression of tetrodotoxin-resistant sodium channels Nav1.8 and Nav1.9 in normal and inflamed rats. Neurosci Lett 355: 45–48. Costigan M, Befort K, Karchewski L, Griffin RS, D’Urso D, Allchorne A, Sitarski J, Mannion JW, Pratt RE, Woolf CJ, 2002. Replicate high-density rat genome oligonucleotide microarrays reveal hundreds of regulated genes in the dorsal root ganglion after peripheral nerve injury. BMC Neurosci 3: 16. Coward K, Aitken A, Powell A, Plumpton C, Birch R, Tate S, Bountra C, Anand P, 2001. Plasticity of TTX-sensitive sodium channels PN1 and brain III in injured human nerves. Neuroreport 12: 495–500. Coward K, Plumpton C, Facer P, Birch R, Carlstedt T, Tate S, Bountra C, Anand P, 2000. Immunolocalization of SNS/PN3 and NaN/SNS2 sodium channels in human pain states. Pain 85: 41–50. Cox JJ, Reimann F, Nicholas AK, Thornton G, Roberts E, Springell K, Karbani G, Jafri H, Mannan J, Raashid Y, Al-Gazali L, Hamamy H, Valente EM, Gorman S, Williams R, McHale DP, Wood JN, Gribble FM, Woods CG, 2006. An SCN9A channelopathy causes congenital inability to experience pain. Nature 444: 894–898. Craner MJ, Damarjian TG, Liu S, Hains BC, Lo AC, Black JA, Newcombe J, Cuzner ML, Waxman SG, 2005. Sodium channels contribute to microglia/macrophage activation and function in EAE and MS. Glia 49: 220–229. Craner MJ, Klein JP, Renganathan M, Black JA, Waxman SG, 2002. Changes of sodium channel expression in experimental painful diabetic neuropathy. Ann Neurol 52: 786–792. Craner MJ, Lo AC, Black JA, Waxman SG, 2003. Abnormal sodium channel distribution in optic nerve axons in a model of inflammatory demyelination. Brain 126: 1552–1561. Cummins TR, Aglieco F, Renganathan M, Herzog RI, Dib-Hajj SD, Waxman SG, 2001. Nav1.3 sodium channels: rapid repriming and slow closed-state inactivation display quantitative differences after expression in a mammalian cell line and in spinal sensory neurons. J Neurosci 21: 5952–5961. Cummins TR, Black JA, Dib-Hajj SD, Waxman SG, 2000. Glial-derived neurotrophic factor upregulates expression of functional SNS and NaN sodium channels and their currents in axotomized dorsal root ganglion neurons. J Neurosci 20: 8754–8761.

REFERENCES

95

Cummins TR, Dib-Hajj SD, Black JA, Akopian AN, Wood JN, Waxman SG, 1999. A novel persistent tetrodotoxin-resistant sodium current in SNS-null and wild-type small primary sensory neurons. J Neurosci 19: RC43. Cummins TR, Dib-Hajj SD, Waxman SG, 2004. Electrophysiological properties of mutant Nav1.7 sodium channels in a painful inherited neuropathy. J Neurosci 24: 8232– 8236. Cummins TR, Howe JR, Waxman SG, 1998. Slow closed-state inactivation: a novel mechanism underlying ramp currents in cells expressing the hNE/PN1 sodium channel. J Neurosci 18: 9607–9619. Cummins TR, Waxman SG, 1997. Downregulation of tetrodotoxin-resistant sodium currents and upregulation of a rapidly repriming tetrodotoxin-sensitive sodium current in small spinal sensory neurons after nerve injury. J Neurosci 17: 3503–3514. Decosterd I, Ji RR, Abdi S, Tate S, Woolf CJ, 2002. The pattern of expression of the voltagegated sodium channels Nav1.8 and Nav1.9 does not change in uninjured primary sensory neurons in experimental neuropathic pain models. Pain 96: 269–277. Decosterd I, Woolf CJ, 2000. Spared nerve injury: an animal model of persistent peripheral neuropathic pain. Pain 87: 149–158. Devor M, 2006. Sodium channels and mechanisms of neuropathic pain. J Pain 7: S3–S12. Devor M, Keller CH, Deerinck TJ, Levinson SR, Ellisman MH, 1989. Naþ channel accumulation on axolemma of afferent endings in nerve end neuromas in Apteronotus. Neurosci Lett 102: 149–154. Dib-Hajj S, Black JA, Cummins TR, Waxman SG, 2002. NaN/Nav1.9: a sodium channel with unique properties. Trends Neurosci 25: 253–259. Dib-Hajj S, Black JA, Felts P, Waxman SG, 1996. Down-regulation of transcripts for Na channel alpha-SNS in spinal sensory neurons following axotomy. Proc Natl Acad Sci USA 93: 14950–14954. Dib-Hajj SD, Black JA, Cummins TR, Kenney AM, Kocsis JD, Waxman SG, 1998a. Rescue of alpha-SNS sodium channel expression in small dorsal root ganglion neurons after axotomy by nerve growth factor in vivo. J Neurophysiol 79: 2668–2676. Dib-Hajj SD, Fjell J, Cummins TR, Zheng Z, Fried K, LaMotte R, Black JA, Waxman SG, 1999. Plasticity of sodium channel expression in DRG neurons in the chronic constriction injury model of neuropathic pain. Pain 83: 591–600. Dib-Hajj SD, Rush AM, Cummins TR, Hisama FM, Novella S, Tyrrell L, Marshall L, Waxman SG, 2005. Gain-of-function mutation in Nav1.7 in familial erythromelalgia induces bursting of sensory neurons. Brain 128: 1847–1854. Dib-Hajj SD, Tyrrell L, Black JA, Waxman SG, 1998b. NaN, a novel voltage-gated Na channel, is expressed preferentially in peripheral sensory neurons and down-regulated after axotomy. Proc Natl Acad Sci USA 95: 8963–8968. Dib-Hajj Sea, 2006. Mutations in the sodium channel Nav1.7 underlie inherited erythromelalgia. Drug Discov Today Dis Mech 3: 343–350. Dietrich PS, McGivern JG, Delgado SG, Koch BD, Eglen RM, Hunter JC, Sangameswaran L, 1998. Functional analysis of a voltage-gated sodium channel and its splice variant from rat dorsal root ganglia. J Neurochem 70: 2262–2272. Diss JK, Stewart D, Pani F, Foster CS, Walker MM, Patel A, Djamgoz MB, 2005. A potential novel marker for human prostate cancer: voltage-gated sodium channel expression in vivo. Prostate Cancer Prostatic Dis 8: 266–273.

96

VOLTAGE-GATED SODIUM CHANNELS

Djouhri L, Fang X, Okuse K, Wood JN, Berry CM, Lawson SN, 2003a. The TTX-resistant sodium channel Nav1.8 (SNS/PN3): expression and correlation with membrane properties in rat nociceptive primary afferent neurons. J Physiol Lond 550: 739–752. Djouhri L, Newton R, Levinson SR, Berry CM, Carruthers B, Lawson SN, 2003b. Sensory and electrophysiological properties of guinea-pig sensory neurones expressing Nav 1.7 (PN1) Naþ channel alpha subunit protein. J Physiol Lond 546: 565–576. Drenth JP, te Morsche RH, Guillet G, Taieb A, Kirby RL, Jansen JB, 2005. SCN9A mutations define primary erythermalgia as a neuropathic disorder of voltage gated sodium channels. J Invest Dermatol 124: 1333–1338. England JD, Happel LT, Kline DG, Gamboni F, Thouron CL, Liu ZP, Levinson SR, 1996a. Sodium channel accumulation in humans with painful neuromas. Neurology 47: 272– 276. England S, Bevan S, Docherty RJ, 1996b. PGE2 modulates the tetrodotoxin-resistant sodium current in neonatal rat dorsal root ganglion neurones via the cyclic AMP-protein kinase A cascade. J Physiol Lond 495 (Pt 2): 429–440. Eriksson J, Jablonski A, Persson AK, Hao JX, Kouya PF, Wiesenfeld-Hallin Z, Xu XJ, Fried K, 2005. Behavioral changes and trigeminal ganglion sodium channel regulation in an orofacial neuropathic pain model. Pain 119: 82–94. Fang X, Djouhri L, Black JA, Dib-Hajj SD, Waxman SG, Lawson SN, 2002. The presence and role of the tetrodotoxin-resistant sodium channel Nav1.9 (NaN) in nociceptive primary afferent neurons. J Neurosci 22: 7425–7433. Fang X, Djouhri L, McMullan S, Berry C, Okuse K, Waxman SG, Lawson SN, 2005. trkA is expressed in nociceptive neurons and influences electrophysiological properties via Nav1.8 expression in rapidly conducting nociceptors. J Neurosci 25: 4868–4878. Felts PA, Yokoyama S, Dib-Hajj S, Black JA, Waxman SG, 1997. Sodium channel alphasubunit mRNAs I, II, III, NaG, Na6 and hNE (PN1): different expression patterns in developing rat nervous system. Brain Res Mol Brain Res 45: 71–82. Fertleman CR, Baker MD, Parker KA, Moffatt S, Elmslie FV, Abrahamsen B, Ostman J, Klugbauer N, Wood JN, Gardiner RM, Rees M, 2006. SCN9A mutations in paroxysmal extreme pain disorder: allelic variants underlie distinct channel defects and phenotypes. Neuron 52: 767–774. Fertleman CR, Ferrie CD, 2006. What’s in a name–familial rectal pain syndrome becomes paroxysmal extreme pain disorder. J Neurol Neurosurg Psychiatry 77: 1294–1295. Filliatreau G, Attal N, Hassig R, Guilbaud G, Desmeules J, DiGiamberardino L, 1994. Timecourse of nociceptive disorders induced by chronic loose ligatures of the rat sciatic nerve and changes of the acetylcholinesterase transport along the ligated nerve. Pain 59: 405– 413. Fitzgerald EM, Okuse K, Wood JN, Dolphin AC, Moss SJ, 1999. cAMP-dependent phosphorylation of the tetrodotoxin-resistant voltage-dependent sodium channel SNS. J Physiol Lond 516: Pt 2 433–446. Fjell J, Cummins TR, Dib-Hajj SD, Fried K, Black JA, Waxman SG, 1999a. Differential role of GDNF and NGF in the maintenance of two TTX-resistant sodium channels in adult DRG neurons. Brain Res Mol Brain Res 67: 267–282. Fjell J, Cummins TR, Fried K, Black JA, Waxman SG, 1999b. In vivo NGF deprivation reduces SNS expression and TTX-R sodium currents in IB4-negative DRG neurons. J Neurophysiol 81: 803–810.

REFERENCES

97

Fjell J, Hjelmstrom P, Hormuzdiar W, Milenkovic M, Aglieco F, Tyrrell L, Dib-Hajj S, Waxman SG, Black JA, 2000. Localization of the tetrodotoxin-resistant sodium channel NaN in nociceptors. Neuroreport 11: 199–202. Fraser SP, Diss JK, Chioni AM, Mycielska ME, Pan H, Yamaci RF, Pani F, Siwy Z, Krasowska M, Grzywna Z, Brackenbury WJ, Theodorou D, Koyuturk M, Kaya H, Battaloglu E, De Bella MT, Slade MJ, Tolhurst R, Palmieri C, Jiang J, Latchman DS, Coombes RC, Djamgoz MB, 2005. Voltage-gated sodium channel expression and potentiation of human breast cancer metastasis. Clin Cancer Res 11: 5381–5389. Fried K, Govrin-Lippmann R, Rosenthal F, Ellisman MH, Devor M, 1991. Ultrastructure of afferent axon endings in a neuroma. J Neurocytol 20: 682–701. George AL Jr, 2005. Inherited disorders of voltage-gated sodium channels. J Clin Invest 115: 1990–1999. Gold MS, Reichling DB, Shuster MJ, Levine JD, 1996. Hyperalgesic agents increase a tetrodotoxin-resistant Naþ current in nociceptors. Proc Natl Acad Sci USA 93: 1108–1112. Gold MS, Weinreich D, Kim CS, Wang R, Treanor J, Porreca F, Lai J, 2003. Redistribution of Nav1.8 in uninjured axons enables neuropathic pain. J Neurosci 23: 158–166. Gold MS, Zhang L, Wrigley DL, Traub RJ, 2002. Prostaglandin E(2) modulates TTX-R I(Na) in rat colonic sensory neurons. J Neurophysiol 88: 1512–1522. Goldin AL, 2002. Evolution of voltage-gated Naþ channels. J Exp Biol 205: 575–584. Goldin AL, Barchi RL, Caldwell JH, Hofmann F, Howe JR, Hunter JC, Kallen RG, Mandel G, Meisler MH, Netter YB, Noda M, Tamkun MM, Waxman SG, Wood JN, Catterall WA, 2000. Nomenclature of voltage-gated sodium channels. Neuron 28: 365–368. Gould HJ 3rd England JD, Soignier RD, Nolan P, Minor LD, Liu ZP, Levinson SR, Paul D, 2004. Ibuprofen blocks changes in Nav1.7 and 1.8 sodium channels associated with complete Freund’s adjuvant-induced inflammation in rat. J Pain 5: 270–280. Gould HJ III Gould TN, England JD, Paul D, Liu ZP, Levinson SR, 2000. A possible role for nerve growth factor in the augmentation of sodium channels in models of chronic pain. Brain Res 854: 19–29. Gould HJ III Gould TN, Paul D, England JD, Liu ZP, Reeb SC, Levinson SR, 1999. Development of inflammatory hypersensitivity and augmentation of sodium channels in rat dorsal root ganglia. Brain Res 824: 296–299. Hains BC, Klein JP, Saab CY, Craner MJ, Black JA, Waxman SG, 2003. Upregulation of sodium channel Nav1.3 and functional involvement in neuronal hyperexcitability associated with central neuropathic pain after spinal cord injury. J Neurosci 23: 8881–8892. Hains BC, Saab CY, Klein JP, Craner MJ, Waxman SG, 2004a. Altered sodium channel expression in second-order spinal sensory neurons contributes to pain after peripheral nerve injury. J Neurosci 24: 4832–4839. Hains BC, Saab CY, Lo AC, Waxman SG, 2004b. Sodium channel blockade with phenytoin protects spinal cord axons, enhances axonal conduction, and improves functional motor recovery after contusion SCI. Exp Neurol 188: 365–377. Hains BC, Saab CY, Waxman SG, 2005. Changes in electrophysiological properties and sodium channel Nav1.3 expression in thalamic neurons after spinal cord injury. Brain 128: 2359– 2371. Hains BC, Saab CY, Waxman SG, 2006. Alterations in burst firing of thalamic VPL neurons and reversal by Nav1.3 antisense after spinal cord injury. J Neurophysiol 95: 3343–3352.

98

VOLTAGE-GATED SODIUM CHANNELS

Han C, Rush AM, Dib-Hajj SD, Li S, Xu Z, Wang Y, Tyrrell L, Wang X, Yang Y, Waxman SG, 2006. Sporadic onset of erythermalgia: a gain-of-function mutation in Nav1.7. Ann Neurol 59: 553–558. Harty TP, Dib-Hajj SD, Tyrrell L, Blackman R, Hisama FM, Rose JB, Waxman SG, 2006. Nav1.7 mutant A863P in erythromelalgia: effects of altered activation and steady-state inactivation on excitability of nociceptive dorsal root ganglion neurons. J Neurosci 26: 12566–12575. Hefti FF, Rosenthal A, Walicke PA, Wyatt S, Vergara G, Shelton DL, Davies AM, 2006. Novel class of pain drugs based on antagonism of NGF. Trends Pharmacol Sci 27: 85–91. Henry MA, Sorensen HJ, Johnson LR, Levinson SR, 2005. Localization of the Nav1.8 sodium channel isoform at nodes of Ranvier in normal human radicular tooth pulp. Neurosci Lett 380: 32–36. Herzog RI, Cummins TR, Ghassemi F, Dib-Hajj SD, Waxman SG, 2003. Distinct repriming and closed-state inactivation kinetics of Nav1.6 and Nav1.7 sodium channels in mouse spinal sensory neurons. J Physiol Lond 551: 741–750. Herzog RI, Cummins TR, Waxman SG, 2001. Persistent TTX-resistant Naþ current affects resting potential and response to depolarization in simulated spinal sensory neurons. J Neurophysiol 86: 1351–1364. Hille B, 2001. Ion Channels of Excitable Membranes, 3rd ed., Sinauer Associates, Inc. Hillsley K, Lin JH, Stanisz A, Grundy D, Aerssens J, Peeters PJ, Moechars D, Coulie B, Stead RH, 2006. Dissecting the role of sodium currents in visceral sensory neurons in a model of chronic hyperexcitability using Nav1.8 and Nav1.9 null mice. J Physiol Lond 576: 257–267. Holm AN, Rich A, Miller SM, Strege P, Ou Y, Gibbons S, Sarr MG, Szurszewski JH, Rae JL, Farrugia G, 2002. Sodium current in human jejunal circular smooth muscle cells. Gastroenterology 122: 178–187. Hong S, Morrow TJ, Paulson PE, Isom LL, Wiley JW, 2004. Early painful diabetic neuropathy is associated with differential changes in tetrodotoxin-sensitive and-resistant sodium channels in dorsal root ganglion neurons in the rat. J Biol Chem 279: 29341–29350. Howe JF, Loeser JD, Calvin WH, 1977. Mechanosensitivity of dorsal root ganglia and chronically injured axons: a physiological basis for the radicular pain of nerve root compression. Pain 3: 25–41. Hu SJ, Song XJ, Greenquist KW, Zhang JM, LaMotte RH, 2001. Protein kinase A modulates spontaneous activity in chronically compressed dorsal root ganglion neurons in the rat. Pain 94: 39–46. Hu SJ, Xing JL, 1998. An experimental model for chronic compression of dorsal root ganglion produced by intervertebral foramen stenosis in the rat. Pain 77: 15–23. Hudmon A, Choi JS, Tyrrell L, Black JA, Rush AM, Waxman SG, Dib-Hajj SD, 2008 Phosphorylation of sodium channel Nav1.8 by p38 mitogen-activated protein kinase increases current density in dorsal root ganglion neurons. J Neurosci 28: 3190–3201. Hunt SP, Mantyh PW, 2001. The molecular dynamics of pain control. Nat Rev Neurosci 2: 83–91. Ikeda H, Stark J, Fischer H, Wagner M, Drdla R, Jager T, Sandkuhler J, 2006. Synaptic amplifier of inflammatory pain in the spinal dorsal horn. Science 312: 1659–1662. Jeong SY, Goto J, Hashida H, Suzuki T, Ogata K, Masuda N, Hirai M, Isahara K, Uchiyama Y, Kanazawa I, 2000. Identification of a novel human voltage-gated sodium channel alpha subunit gene, SCN12A. Biochem Biophys Res Commun 267: 262–270. Ji RR, 2004. Peripheral and central mechanisms of inflammatory pain, with emphasis on MAP kinases. Curr Drug Targets Inflamm Allergy 3: 299–303.

REFERENCES

99

Jin X, Gereau RW 2006. Acute p38-mediated modulation of tetrodotoxin-resistant sodium channels in mouse sensory neurons by tumor necrosis factor-alpha. J Neurosci 26: 246–255. Jo T, Nagata T, Iida H, Imuta H, Iwasawa K, Ma J, Hara K, Omata M, Nagai R, Takizawa H, Nagase T, Nakajima T, 2004. Voltage-gated sodium channel expressed in cultured human smooth muscle cells: involvement of SCN9A. FEBS Lett 567: 339–343. Joshi SK, Mikusa JP, Hernandez G, Baker S, Shieh CC, Neelands T, Zhang XF, Niforatos W, Kage K, Han P, Krafte D, Faltynek C, Sullivan JP, Jarvis MF, Honore P, 2006. Involvement of the TTX-resistant sodium channel Nav1.8 in inflammatory and neuropathic, but not postoperative, pain states. Pain 123: 75–82. Kajander KC, Wakisaka S, Bennett GJ, 1992. Spontaneous discharge originates in the dorsal root ganglion at the onset of a painful peripheral neuropathy in the rat. Neurosci Lett 138: 225–228. Kaplan MR, Cho MH, Ullian EM, Isom LL, Levinson SR, Barres BA, 2001. Differential control of clustering of the sodium channels Nav1.2 and Nav1.6 at developing CNS nodes of Ranvier. Neuron 30: 105–119. Kazarinova-Noyes K, Malhotra JD, McEwen DP, Mattei LN, Berglund EO, Ranscht B, Levinson SR, Schachner M, Shrager P, Isom LL, Xiao ZC, 2001. Contactin associates with Naþ channels and increases their functional expression. J Neurosci 21: 7517–7525. Kerr BJ, Souslova V, McMahon SB, Wood JN, 2001. A role for the TTX-resistant sodium channel Nav1.8 in NGF-induced hyperalgesia, but not neuropathic pain. Neuroreport 12: 3077–3080. Khasar SG, Gold MS, Levine JD, 1998. A tetrodotoxin-resistant sodium current mediates inflammatory pain in the rat. Neurosci Lett 256: 17–20. Kim CH, Oh Y, Chung JM, Chung K, 2001. The changes in expression of three subtypes of TTX sensitive sodium channels in sensory neurons after spinal nerve ligation. Brain Res Mol Brain Res 95: 153–161. Kim CH, Oh Y, Chung JM, Chung K, 2002. Changes in three subtypes of tetrodotoxin sensitive sodium channel expression in the axotomized dorsal root ganglion in the rat. Neurosci Lett 323: 125–128. Kim SH, Chung JM, 1992. An experimental model for peripheral neuropathy produced by segmental spinal nerve ligation in the rat. Pain 50: 355–363. Klugbauer N, Lacinova L, Flockerzi V, Hofmann F, 1995. Structure and functional expression of a new member of the tetrodotoxin-sensitive voltage-activated sodium channel family from human neuroendocrine cells. EMBO J 14: 1084–1090. Kral MG, Xiong Z, Study RE, 1999. Alteration of Naþ currents in dorsal root ganglion neurons from rats with a painful neuropathy. Pain 81: 15–24. Kretschmer T, Happel LT, England JD, Nguyen DH, Tiel RL, Beuerman RW, Kline DG, 2002. Accumulation of PN1 and PN3 sodium channels in painful human neuroma-evidence from immunocytochemistry. Acta Neurochir (Wien) 144: 803–810; discussion 810. Kretschmer T, Nguyen DH, Beuerman RW, Tiel RL, Kline DG, 2004. Elevated ankyrin G in a plexiform neurofibroma and neuromas associated with pain. J Clin Neurosci 11: 886–889. Lai J, Gold MS, Kim CS, Bian D, Ossipov MH, Hunter JC, Porreca F, 2002. Inhibition of neuropathic pain by decreased expression of the tetrodotoxin-resistant sodium channel, Nav1.8. Pain 95: 143–152. Lai J, Porreca F, Hunter JC, Gold MS, 2004. Voltage-gated sodium channels and hyperalgesia. Annu Rev Pharmacol Toxicol 44: 371–397.

100

VOLTAGE-GATED SODIUM CHANNELS

Laird JM, Souslova V, Wood JN, Cervero F, 2002. Deficits in visceral pain and referred hyperalgesia in Nav1.8 (SNS/PN3)-null mice. J Neurosci 22: 8352–8356. Lampert A, Dib-Hajj SD, Tyrrell L, Waxman SG, 2006. Size matters: Erythromelalgia mutation S241T in Nav1.7 alters channel gating. J Biol Chem 281: 36029–36035. Leffler A, Cummins TR, Dib-Hajj SD, Hormuzdiar WN, Black JA, Waxman SG, 2002. GDNF and NGF reverse changes in repriming of TTX-sensitive Naþ currents following axotomy of dorsal root ganglion neurons. J Neurophysiol 88: 650–658. Lewin-Kowalik J, Marcol W, Kotulska K, Mandera M, Klimczak A, 2006. Prevention and management of painful neuroma. Neurol Med Chir (Tokyo) 46: 62–67 discussion 67–68. Lindia JA, Kohler MG, Martin WJ, Abbadie C, 2005. Relationship between sodium channel Nav1.3 expression and neuropathic pain behavior in rats. Pain 117: 145–153. Liu C, Cummins TR, Tyrrell L, Black JA, Waxman SG, Dib-Hajj SD, 2005. CAP-1A is a novel linker that binds clathrin and the voltage-gated sodium channel Nav1.8. Mol Cell Neurosci 28: 636–649. Liu X, Chung K, Chung JM, 1999. Ectopic discharges and adrenergic sensitivity of sensory neurons after spinal nerve injury. Brain Res 849: 244–247. Liu X, Zhou JL, Chung K, Chung JM, 2001. Ion channels associated with the ectopic discharges generated after segmental spinal nerve injury in the rat. Brain Res 900: 119–127. Ma C, LaMotte RH, 2005. Enhanced excitability of dissociated primary sensory neurons after chronic compression of the dorsal root ganglion in the rat. Pain 113: 106–112. Ma W, Bisby MA, 1998. Increase of preprotachykinin mRNA and substance P immunoreactivity in spared dorsal root ganglion neurons following partial sciatic nerve injury. Eur J Neurosci 10: 2388–2399. Maier SK, Westenbroek RE, Schenkman KA, Feigl EO, Scheuer T, Catterall WA, 2002. An unexpected role for brain-type sodium channels in coupling of cell surface depolarization to contraction in the heart. Proc Natl Acad Sci USA 99: 4073–4078. Malykhina AP, Qin C, Foreman RD, Akbarali HI, 2004. Colonic inflammation increases Naþ currents in bladder sensory neurons. Neuroreport 15: 2601–2605. Matzner O, Devor M, 1994. Hyperexcitability at sites of nerve injury depends on voltagesensitive Naþ channels. J Neurophysiol 72: 349–359. Meisler MH, Kearney JA, 2005. Sodium channel mutations in epilepsy and other neurological disorders. J Clin Invest 115: 2010–2017. Michiels JJ, te Morsche RH, Jansen JB, Drenth JP, 2005. Autosomal dominant erythermalgia associated with a novel mutation in the voltage-gated sodium channel alpha subunit Nav1.7. Arch Neurol 62: 1587–1590. Moss AJ, Kass RS, 2005. Long QT syndrome: from channels to cardiac arrhythmias. J Clin Invest 115: 2018–2024. Myers RR, Heckman HM, Rodriguez M, 1996. Reduced hyperalgesia in nerve-injured WLD mice: relationship to nerve fiber phagocytosis, axonal degeneration, and regeneration in normal mice. Exp Neurol 141: 94–101. Nassar MA, Baker MD, Levato A, Ingram R, Mallucci G, McMahon SB, Wood JN, 2006. Nerve injury induces robust allodynia and ectopic discharges in Nav1.3 null mutant mice. Mol Pain 2: 33. Nassar MA, Levato A, Stirling LC, Wood JN, 2005. Neuropathic pain develops normally in mice lacking both Nav1.7 and Nav1.8. Mol Pain 1: 24.

REFERENCES

101

Nassar MA, Stirling LC, Forlani G, Baker MD, Matthews EA, Dickenson AH, Wood JN, 2004. Nociceptor-specific gene deletion reveals a major role for Nav1.7 (PN1) in acute and inflammatory pain. Proc Natl Acad Sci USA 101: 12706–12711. Novakovic SD, Tzoumaka E, McGivern JG, Haraguchi M, Sangameswaran L, Gogas KR, Eglen RM, Hunter JC, 1998. Distribution of the tetrodotoxin-resistant sodium channel PN3 in rat sensory neurons in normal and neuropathic conditions. J Neurosci 18: 2174–2187. Novella SP, Hisama FM, Dib-Hajj SD, Waxman SG, 2007. A case of inherited erythromelalgia. Nat Clin Pract Neurol 3: 229–234. Okuse K, Chaplan SR, McMahon SB, Luo ZD, Calcutt NA, Scott BP, Akopian AN, Wood JN, 1997. Regulation of expression of the sensory neuron-specific sodium channel SNS in inflammatory and neuropathic pain. Mol Cell Neurosci 10: 196–207. Petersen M, Zhang J, Zhang JM, LaMotte RH, 1996. Abnormal spontaneous activity and responses to norepinephrine in dissociated dorsal root ganglion cells after chronic nerve constriction. Pain 67: 391–397. Pierre K, Dupouy B, Allard M, Poulain DA, Theodosis DT, 2001. Mobilization of the cell adhesion glycoprotein F3/contactin to axonal surfaces is activity dependent. Eur J Neurosci 14: 645–656. Porreca F, Lai J, Bian D, Wegert S, Ossipov MH, Eglen RM, Kassotakis L, Novakovic S, Rabert DK, Sangameswaran L, Hunter JC, 1999. A comparison of the potential role of the tetrodotoxin-insensitive sodium channels, PN3/SNS and NaN/SNS2, in rat models of chronic pain. Proc Natl Acad Sci USA 96: 7640–7644. Priest BT, Murphy BA, Lindia JA, Diaz C, Abbadie C, Ritter AM, Liberator P, Iyer LM, Kash SF, Kohler MG, Kaczorowski GJ, MacIntyre DE, Martin WJ, 2005. Contribution of the tetrodotoxin-resistant voltage-gated sodium channel Nav1.9 to sensory transmission and nociceptive behavior. Proc Natl Acad Sci USA 102: 9382–9387. Renganathan M, Cummins TR, Waxman SG, 2001. Contribution of Nav1.8 sodium channels to action potential electrogenesis in DRG neurons. J Neurophysiol 86: 629–640. Rice AS, Hill RG, 2006. New treatments for neuropathic pain. Annu Rev Med 57: 535–551. Rizzo MA, Kocsis JD, Waxman SG, 1994. Slow sodium conductances of dorsal root ganglion neurons: intraneuronal homogeneity and interneuronal heterogeneity. J Neurophysiol 72: 2796–2815. Rizzo MA, Kocsis JD, Waxman SG, 1995. Selective loss of slow and enhancement of fast Naþ currents in cutaneous afferent dorsal root ganglion neurones following axotomy. Neurobiol Dis 2: 87–96. Roza C, Laird JM, Souslova V, Wood JN, Cervero F, 2003. The tetrodotoxin-resistant Naþ channel Nav1.8 is essential for the expression of spontaneous activity in damaged sensory axons of mice. J Physiol Lond 550: 921–926. Rugiero F, Mistry M, Sage D, Black JA, Waxman SG, Crest M, Clerc N, Delmas P, Gola M, 2003. Selective expression of a persistent tetrodotoxin-resistant Naþ current and Nav1.9 subunit in myenteric sensory neurons. J Neurosci 23: 2715–2725. Rush AM, Craner MJ, Kageyama T, Dib-Hajj SD, Waxman SG, Ranscht B, 2005. Contactin regulates the current density and axonal expression of tetrodotoxin-resistant but not tetrodotoxin-sensitive sodium channels in DRG neurons. Eur J Neurosci 22: 39–49. Rush AM, Cummins TR, Waxman SG, 2007. Multiple sodium channels and their roles in electrogenesis within dorsal root ganglion neurons. J Physiol Lond 579: 1–14.

102

VOLTAGE-GATED SODIUM CHANNELS

Rush AM, Dib-Hajj SD, Liu S, Cummins TR, Black JA, Waxman SG, 2006. A single sodium channel mutation produces hyper- or hypoexcitability in different types of neurons. Proc Natl Acad Sci USA 103: 8245–8250. Rush AM, Waxman SG, 2004. PGE2 increases the tetrodotoxin-resistant Nav1.9 sodium current in mouse DRG neurons via G-proteins. Brain Res 1023: 264–271. Saleh S, Yeung SY, Prestwich S, Pucovsky V, Greenwood I, 2005. Electrophysiological and molecular identification of voltage-gated sodium channels in murine vascular myocytes. J Physiol Lond 568: 155–169. Sangameswaran L, Delgado SG, Fish LM, Koch BD, Jakeman LB, Stewart GR, Sze P, Hunter JC, Eglen RM, Herman RC, 1996. Structure and function of a novel voltage-gated, tetrodotoxinresistant sodium channel specific to sensory neurons. J Biol Chem 271: 5953–5956. Sangameswaran L, Fish LM, Koch BD, Rabert DK, Delgado SG, Ilnicka M, Jakeman LB, Novakovic S, Wong K, Sze P, Tzoumaka E, Stewart GR, Herman RC, Chan H, Eglen RM, Hunter JC, 1997. A novel tetrodotoxin-sensitive, voltage-gated sodium channel expressed in rat and human dorsal root ganglia. J Biol Chem 272: 14805–14809. Sasaki M, Black JA, Lankford KL, Tokuno HA, Waxman SG, Kocsis JD, 2006. Molecular reconstruction of nodes of Ranvier after remyelination by transplanted olfactory ensheathing cells in the demyelinated spinal cord. J Neurosci 26: 1803–1812. Scadding JW, 1981. Development of ongoing activity, mechanosensitivity, and adrenaline sensitivity in severed peripheral nerve axons. Exp Neurol 73: 345–364. Schafer DP, Custer AW, Shrager P, Rasband MN, 2006. Early events in node of Ranvier formation during myelination and remyelination in the PNS. Neuron Glia Biol 2: 69–79. Schaller KL, Caldwell JH, 2000. Developmental and regional expression of sodium channel isoform NaCh6 in the rat central nervous system. J Comp Neurol 420: 84–97. Schaller KL, Krzemien DM, Yarowsky PJ, Krueger BK, Caldwell JH, 1995. A novel, abundant sodium channel expressed in neurons and glia. J Neurosci 15: 3231–3242. Shah BS, Rush AM, Liu S, Tyrrell L, Black JA, Dib-Hajj SD, Waxman SG, 2004. Contactin associates with sodium channel Nav1.3 in native tissues and increases channel density at the cell surface. J Neurosci 24: 7387–7399. Sleeper AA, Cummins TR, Dib-Hajj SD, Hormuzdiar W, Tyrrell L, Waxman SG, Black JA, 2000. Changes in expression of two tetrodotoxin-resistant sodium channels and their currents in dorsal root ganglion neurons after sciatic nerve injury but not rhizotomy. J Neurosci 20: 7279–7289. Sommer C, Schafers M, 1998. Painful mononeuropathy in C57BL/Wld mice with delayed wallerian degeneration: differential effects of cytokine production and nerve regeneration on thermal and mechanical hypersensitivity. Brain Res 784: 154–162. Song XJ, Zhang JM, Hu SJ, LaMotte RH, 2003. Somata of nerve-injured sensory neurons exhibit enhanced responses to inflammatory mediators. Pain 104: 701–709. Study RE, Kral MG, 1996. Spontaneous action potential activity in isolated dorsal root ganglion neurons from rats with a painful neuropathy. Pain 65: 235–242. Sun JH, Yang B, Donnelly DF, Ma C, LaMotte RH, 2006. MCP-1 enhances excitability of nociceptive neurons in chronically compressed dorsal root ganglia. J Neurophysiol 96: 2189–2199. Tan ZY, Donnelly DF, LaMotte RH, 2006. Effects of a chronic compression of the dorsal root ganglion on voltage-gated Naþ and Kþ currents in cutaneous afferent neurons. J Neurophysiol 95: 1115–1123.

REFERENCES

103

Tanaka M, Cummins TR, Ishikawa K, Dib-Hajj SD, Black JA, Waxman SG, 1998. SNS Naþ channel expression increases in dorsal root ganglion neurons in the carrageenan inflammatory pain model. Neuroreport 9: 967–972. Tate S, Benn S, Hick C, Trezise D, John V, Mannion RJ, Costigan M, Plumpton C, Grose D, Gladwell Z, Kendall G, Dale K, Bountra C, Woolf CJ, 1998. Two sodium channels contribute to the TTX-R sodium current in primary sensory neurons. Nat Neurosci 1: 653–655. Toledo-Aral JJ, Brehm P, Halegoua S, Mandel G, 1995. A single pulse of nerve growth factor triggers long-term neuronal excitability through sodium channel gene induction. Neuron 14: 607–611. Toledo-Aral JJ, Moss BL, He ZJ, Koszowski AG, Whisenand T, Levinson SR, Wolf JJ, SilosSantiago I, Halegoua S, Mandel G, 1997. Identification of PN1, a predominant voltagedependent sodium channel expressed principally in peripheral neurons. Proc Natl Acad Sci USA 94: 1527–1532. Van Wart A, Boiko T, Trimmer JS, Matthews G, 2005. Novel clustering of sodium channel Nav1.1 with ankyrin-G and neurofascin at discrete sites in the inner plexiform layer of the retina. Mol Cell Neurosci 28: 661–673. Van Zundert J, Harney D, Joosten EA, Durieux ME, Patijn J, Prins MH, Van Kleef M, 2006. The role of the dorsal root ganglion in cervical radicular pain: diagnosis, pathophysiology, and rationale for treatment. Reg Anesth Pain Med 31: 152–167. Venance SL, Cannon SC, Fialho D, Fontaine B, Hanna MG, Ptacek LJ, Tristani-Firouzi M, Tawil R, Griggs RC, 2006. The primary periodic paralyses: diagnosis, pathogenesis and treatment. Brain 129: 8–17. Vernadakis AJ, Koch H, Mackinnon SE, 2003. Management of neuromas. Clin Plast Surg 30: 247–268 vii. Vijayaragavan K, Boutjdir M, Chahine M, 2004. Modulation of Nav1.7 and Nav1.8 peripheral nerve sodium channels by protein kinase A and protein kinase C. J Neurophysiol 91: 1556– 1569. Viswanathan PC, Balser JR, 2004. Inherited sodium channelopathies: a continuum of channel dysfunction. Trends Cardiovasc Med 14: 28–35. Waxman SG, 1999. The molecular pathophysiology of pain: abnormal expression of sodium channel genes and its contributions to hyperexcitability of primary sensory neurons. Pain Suppl 6: S133–S140. Waxman SG, 2001. Transcriptional channelopathies: an emerging class of disorders. Nat Rev Neurosci 2: 652–659. Waxman SG, Craner MJ, Black JA, 2004. Naþ channel expression along axons in multiple sclerosis and its models. Trends Pharmacol Sci 25: 584–591. Waxman SG, Dib-Hajj S, 2005. Erythermalgia: molecular basis for an inherited pain syndrome. Trends Mol Med 11: 555–562. Waxman SG, Dib-Hajj S, Cummins TR, Black JA, 1999. Sodium channels and pain. Proc Natl Acad Sci USA 96: 7635–7639. Waxman SG, Hains BC, 2006. Fire and phantoms after spinal cord injury: Naþ channels and central pain. Trends Neurosci 29: 207–215. Waxman SG, Kocsis JD, Black JA, 1994. Type III sodium channel mRNA is expressed in embryonic but not adult spinal sensory neurons, and is reexpressed following axotomy. J Neurophysiol 72: 466–470.

104

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Westenbroek RE, Merrick DK, Catterall WA, 1989. Differential subcellular localization of the RI and RII Naþ channel subtypes in central neurons. Neuron 3: 695–704. Westenbroek RE, Noebels JL, Catterall WA, 1992. Elevated expression of type II Naþ channels in hypomyelinated axons of shiverer mouse brain. J Neurosci 12: 2259–2267. Whitaker W, Faull R, Waldvogel H, Plumpton C, Burbidge S, Emson P, Clare J, 1999. Localization of the type VI voltage-gated sodium channel protein in human CNS. Neuroreport 10: 3703–3709. Whitaker WR, Faull RL, Waldvogel HJ, Plumpton CJ, Emson PC, Clare JJ, 2001. Comparative distribution of voltage-gated sodium channel proteins in human brain. Brain Res Mol Brain Res 88: 37–53. Wood JN, Boorman JP, Okuse K, Baker MD, 2004. Voltage-gated sodium channels and pain pathways. J Neurobiol 61: 55–71. Woolf CJ, 2004. Dissecting out mechanisms responsible for peripheral neuropathic pain: implications for diagnosis and therapy. Life Sci 74: 2605–2610. Xiao HS, Huang QH, Zhang FX, Bao L, Lu YJ, Guo C, Yang L, Huang WJ, Fu G, Xu SH, Cheng XP, Yan Q, Zhu ZD, Zhang X, Chen Z, Han ZG, Zhang X, 2002. Identification of gene expression profile of dorsal root ganglion in the rat peripheral axotomy model of neuropathic pain. Proc Natl Acad Sci USA 99: 8360–8365. Xie Y, Zhang J, Petersen M, LaMotte RH, 1995. Functional changes in dorsal root ganglion cells after chronic nerve constriction in the rat. J Neurophysiol 73: 1811–1820. Yang L, Zhang FX, Huang F, Lu YJ, Li GD, Bao L, Xiao HS, Zhang X, 2004a. Peripheral nerve injury induces trans-synaptic modification of channels, receptors and signal pathways in rat dorsal spinal cord. Eur J Neurosci 19: 871–883. Yang Y, Wang Y, Li S, Xu Z, Li H, Ma L, Fan J, Bu D, Liu B, Fan Z, Wu G, Jin J, Ding B, Zhu X, Shen Y, 2004b. Mutations in SCN9A, encoding a sodium channel alpha subunit, in patients with primary erythermalgia. J Med Genet 41: 171–174. Yeomans DC, Levinson SR, Peters MC, Koszowski AG, Tzabazis AZ, Gilly WF, Wilson SP, 2005. Decrease in inflammatory hyperalgesia by herpes vector-mediated knockdown of Nav1.7 sodium channels in primary afferents. Hum Gene Ther 16: 271–277. Yoshimura N, Seki S, Novakovic SD, Tzoumaka E, Erickson VL, Erickson KA, Chancellor MB, de Groat WC, 2001. The involvement of the tetrodotoxin-resistant sodium channel Nav1.8 (PN3/SNS) in a rat model of visceral pain. J Neurosci 21: 8690–8696. Zhang JM, Song XJ, LaMotte RH, 1997. An in vitro study of ectopic discharge generation and adrenergic sensitivity in the intact, nerve-injured rat dorsal root ganglion. Pain 72: 51–57. Zhang JM, Song XJ, LaMotte RH, 1999. Enhanced excitability of sensory neurons in rats with cutaneous hyperalgesia produced by chronic compression of the dorsal root ganglion. J Neurophysiol 82: 3359–3366. Zhang YH, Vasko MR, Nicol GD, 2002. Ceramide, a putative second messenger for nerve growth factor, modulates the TTX-resistant Naþ current and delayed rectifier Kþ current in rat sensory neurons. J Physiol Lond 544: 385–402. Zhou Z, Davar G, Strichartz G, 2002. Endothelin-1 (ET-1) selectively enhances the activation gating of slowly inactivating tetrodotoxin-resistant sodium currents in rat sensory neurons: a mechanism for the pain-inducing actions of ET-1. J Neurosci 22: 6325–6330.

4 THE ROLE OF ION CHANNELS IN THE ETIOLOGY AND DEVELOPMENT OF GLIOMAS Amy K. Weaver and Harald Sontheimer Department of Neurobiology and Center for Glial Biology in Medicine, The University of Alabama, Birmingham, AL 35294, USA

4.1 INTRODUCTION Over the last decades, several studies have characterized ion channel expression in glial cells (Sontheimer and Waxman, 1993; Ransom and Sontheimer, 1995; Sontheimer et al., 1996; Bordey and Sontheimer, 1997, 1998b, 2000; MacFarlane and Sontheimer, 1997, 2000a; Parkerson and Sontheimer, 2004; Olsen et al., 2006). By and large, these studies agree on the notion that these channels do not participate in any form of electrical signaling but instead engage in extracellular ion homeostasis or support fundamental aspects of cell biology. An excellent example for a homeostatic role of ion channels is the glial inwardly rectifying Kþ channel, Kir4.1, which has been shown to constitute the primary uptake pathway for neuronally released Kþ from the extracellular space (Sontheimer et al., 1996; Higashi et al., 2001; Olsen et al., 2006, 2007). Its elimination results in a complete breakdown of extracellular Kþ homeostasis (Neusch et al., 2006; Kucheryavykh et al., 2007). Numerous studies have demonstrated a role for delayed rectifying Kþ channels of the Kv1.x family in cell cycle progression. In astrocytes, Kv1.5 is reversibly phosphorylated by Src kinase as cells enter S-phase leading to a marked cell depolarization. Suppression of Kv1.5 activity or inhibition of Src kinases causes reversible cell cycle arrest (MacFarlane and Sontheimer, 2000b). Importantly, acute Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

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injury causes gliosis and a similar activation of Kv1.5 in many glial cell types, including retinal glial cells (Bordey et al., 2001). As the participation of ion channels in cell proliferation became more compelling, numerous investigations examined tumor cells that show aberrant, uncontrolled cell proliferation. These studies propose that certain channels, the ether-a-go-go (EAG) channel family being a prominent example, may be so strongly connected to the oncogenic phenotype of these cancers that these channels could actually be considered oncogenes (Meyer et al., 1999; Kunzelmann, 2005; Camacho, 2006). This chapter will focus exclusively on a subset of primary central nervous system (CNS) tumors, gliomas, to illustrate how ion channel function promotes the unique biology of these cancers. Most of the primary brain tumors originate from glial cells or their progenitor cells and are collectively called gliomas. These rapidly proliferating tumors include astrocytomas, glioblastomas, and oligodendrogliomas and share an unusual propensity to invade the brain. This feature is particularly pronounced in the highest tumor grade, glioblastoma multiforme, which is the most frequent adult glioma. By invading the normal brain parenchyma, they escape complete surgical resection and make effective radiation therapy challenging. Consequently, gliomas carry an unusually high mortality rate with most patients succumbing to the disease within 12–18 months of diagnosis, typically due to regrowth of the tumor (Kleihues et al., 1995; Ohgaki and Kleihues, 2005). Glioma migration/invasion requires complex and well-arranged interactions of molecular motors with the cytoskeleton and adhesion sites of cells interacting with other cells or the extracellular matrix surrounding them (for reviews see Giese et al., 1994; Gladson, 1999; Uhm et al., 1999; Tonn and Goldbrunner, 2003; Visted et al., 2003; Demuth and Berens, 2004; Zamecnik, 2005). It also involves the production and release of matrix-degrading enzymes (Ohnishi et al., 1993; Maidment et al., 1996; Deryugina et al., 1997; Treasurywala and Berens, 1998; Belien et al., 1999; Gladson, 1999; Nakada et al., 2003) and chemotactic interactions with neighboring cells (Merzak et al., 1994; Giese et al., 1995, 1998; Chicoine and Silbergeld, 1997; Maidment et al., 1997; Manning et al., 2000; Ritch et al., 2003; Farin et al., 2006). Many of these features are shared with other cancers. However, in stark contrast to other cancers, glioma cells spread by active cell migration rather than spreading passively via hematogenous routes. As a result, gliomas rarely ever metastasize outside the CNS, they readily invade and metastasize within the brain and spinal cord. This is somewhat surprising since the extracellular space in the mature brain is very small. Cells that navigate the narrow and tortuous extracellular space will need to be able to adjust their shape and volume to fit through these narrow spaces. Indeed, as illustrated for two examples in Fig. 4.1, glioma cells that penetrate through the extracellular space are elongated, wedge-shaped with slender cell processes (Soroceanu et al., 1999). This morphology is consistent with an overall shrinkage of the cell body and an elongation of invadipodia, the leading processes of cells actively engaged in invasion. Cell shrinkage requires the secretion of cytoplasm, principally KCl and water from the cell. As further discussed below, and illustrated in Fig. 4.6a, this is accomplished by the coordinated activity of defined Kþ and Cl
channels, which are the principal diffusional release pathways for these ions.

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FIGURE 4.1 (a) Semithin section through a coculture of tumor spheroids and fetal rat brain aggregates, stained with toluidine blue. Glioma cells are seen advancing between two normal rat brain cells (arrows). Scale bar, 20 mm. (b) Area of detail is of the same preparation as in (a), analyzed by transmission electron microcopy. Glioma cells are easily recognized because of the abundance of ribosomes and other organelles that incorporate lead citrate and give a darker appearance. Arrows indicate area of contact between an elongated tumor cell and two other membranes, presumably of the fetal rat brain. Scale bar, 1 mm. (Adapted from Soroceanu et al., 1999).

4.2 Kþ CHANNELS IN GLIOMAS Potassium channels are essential to establish and maintain a negative resting potential in many cells including nonexcitable cells. At least 40 different Kþ channels can now be identified molecularly and many cells express more than one channel type. In the earliest biophysical studies of glial cells (Kuffler, 1967), it was noted that the glial cell membrane appeared to be almost exclusively permeable to Kþ ions. Not surprisingly, later studies showed that glial cells express Kþ channels at a very high density. Through the use of genetic tools, it has now become evident that the resting potential of astrocytes and retinal glial cells is mainly established by the activity of Kir4.1, a voltage-dependent inwardly rectifying Kþ channel (Chvatal et al., 1995; Li et al., 2001; Kofuji et al., 2002). Kir4.1 channel expression arising upon differentiation of progenitor cells into mature postmitotic cells is developmentally regulated in astrocytes and in oligodendrocytes (Ransom and Sontheimer, 1995). The malignant transformation of astrocytes, that is, formation of gliomas, correlates with a depolarization of the resting membrane potential and, indeed, this seems to be due to the loss of functional Kir4.1 channels (Olsen and Sontheimer, 2004). Surprisingly however, Kir4.1 channels are still expressed, but are mislocalized to a perinuclear region, presumably the ER indicating an inability to correctly traffic the channel to the membrane. Recombinant expression studies suggest that insertion of Kir4.1 into glioma membranes reestablishes a negative resting potential and is sufficient to cause growth inhibition (Higashimori and Sontheimer, 2007). This study also demonstrates that a downregulation of Kir4.1 channels is required to allow glial and glioma cells to

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proliferate and from a mechanistic point of view, a depolarizing shift in resting membrane potential is essential for a cell’s progression through S-phase. An involvement of Kþ channels in growth control of cancer cells is now widely accepted (for review see Wonderlin and Strobl, 1996; Kunzelmann, 2005), suggesting that Kþ channels may be excellent therapeutic cancer targets. Many studies have implicated, in particular, two Kþ channel families in growth control. These are the large conductance, calcium-activated Kþ (BK) channels and the human ether-agogo-like (hERG) channels. While very little is known about hERG channels in the glioma, BK channels are highly expressed and are probably the most well-studied ion channel in the glioma field. A decade ago we noted the presence of large outward currents in cells from patient biopsies that were activated by membrane depolarizations larger than þ50 mV (Ullrich et al., 1998). Subsequent studies identified these currents as Kþ selective, sensitive to micromolar concentrations of Ca2þ, and inhibited by Iberiotoxin (IbTX), tetraethylammonium(TEA) chloride, and tetrandrine (Ransom and Sontheimer, 2001), all indicative of BK channel expression and representative examples are shown in Fig. 4.2a. The BK channels are a member of the family of calcium-activated Kþ channels and their names derive from their very large single-channel conductances (>200 pS in glioma cells). These channels are unique among Kþ channels in that their activity can be modulated by changes in intracellular calcium concentration in addition to changes in membrane voltage. Interestingly, the BK channel expressed in glioma cells is a novel splice variant with a 36 amino acid insert that confers an increased sensitivity to calcium when compared to other known BK channel splice variants (Ransom et al., 2002; Liu et al., 2002) (Fig. 4.2b). Because of this, these channels are well suited to respond to various mitogenic and motogenic signals that affect intracellular calcium concentration. This combined with the fact that BK currents account for most of the voltage-dependent currents in glioma cells has led investigators to examine the function of these channels in gliomas. Through the use of specific pharmacology, it has been discovered that BK channels indeed play a role in glioma cell proliferation, albeit a complicated one. Straightforward examination of glioma cell proliferation under normal culture conditions has revealed no effect of the specific BK channel inhibitors, IbTX and paxilline. However, when serum was removed from the culture media (Weaver et al., 2004) or when cells were stimulated to proliferate by raising extracellular Kþ concentration (Basrai et al., 2002), these drugs inhibited glioma cell proliferation. While this may seem counterintuitive, it actually may reveal a significant problem with the use of glioma cell lines in research. The brain does not exist in a state of unlimited nutrient and oxygen supply. In fact, gliomas proliferate at such a rate that they rapidly exhaust the nutrients and oxygen that are available to them, suggesting that current culture conditions containing fetal calf serum and 21 % oxygen do not accurately mimic the conditions in vivo. In agreement with this, gliomas are characterized by overexpression and/or constitutive activation of growth factor receptors (Collins, 1995; Di et al., 2000). One such receptor, the erbB2 growth factor receptor, is constitutively active in gliomas and appears to be capable of modulating BK channel activity through changes

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FIGURE 4.2 BK currents in glioma cells (a) whole-cell currents from a D54-MG cell with the indicated concentrations of iberiotoxin, a selective BK channel inhibitor. Inset: The time course of current inhibition in this cell by iberiotoxin (current measured at þ140 mV). Iberiotoxin inhibition was reversible. (b) Average activation curves with the indicated [Ca2þ]i. Data are from 4–17 outside-out patches from D54MG glioma cells. Error bars are omitted for clarity. Solid lines are drawn according to the Boltzmann equation. (Adapted from Liu et al., 2002; Ransom et al., 2002).

in [Ca2þ]i (Olsen et al., 2005) providing direct evidence that BK channels can be regulated by proliferative signaling. In addition to its role in proliferation, BK channel involvement in migration/ invasion has also been documented. The results of these studies, however, were also complex. In 2000, Bordey et al. (2000) showed that muscarinic activation of acetycholine (ACh) receptors activated BK channels as well as inhibited glioma cell migration in a two-dimensional motility assay. These results were corroborated 3 years later by Kraft et al. (2003) who noticed that activation of BK channels with phloretin and NS1619, a synthetic BK channel opener, also inhibited migration of another glioma cell line. Contrary to these findings, a later study by Weaver et al. (2006)

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examined the effect of BK channel inhibitors and NS1619 on glioma cell migration across a transwell barrier, a technique used to mimic the physical constraints encountered in vivo, and found that inhibition, not activation, of BK channels inhibited migration of yet another glioma cell. It remains to be seen whether these different effects are due to different assay systems or due to the use of different glioma cell lines. However, a recent study suggests that BK channels are found in lipid rafts on the lamellipodia, the invading edge, of glioma cells where they specifically respond to both mitogenic and motogenic signals (Weaver et al., 2007). Together these findings may indicate that BK channel activation can have different cellular consequences, depending on the upstream signal responsible for the calcium changes activating the channels. In light of the high expression of BK channels in gliomas in vitro and in vivo and their described role in growth control and cell invasion, one may wish to think of them as potential therapeutic targets. Indeed, BK channels have two specific blockers and several synthetic openers making it possible to target these channels without nonspecifically affecting other ion channels. Unfortunately, however, BK channels have fundamental roles in smooth muscle contraction/relaxation and help control bursting frequency in neurons. All of this must be taken into account when designing possible therapeutic strategies to target these channels. The human ether-a-go-go family of ion channels is made up of three subfamilies, the ether-a-go-go or EAG channels, the ether-a-go-go-related or ERK and the ethera-go-go-like or ELK channels. For the purposes of the chapter, we will collectively refer to these channels as the EAG channels. All EAG channels give rise to outwardly rectifying, slowly activating, noninactivating Kþ currents that activate in response to depolarizing voltages. This family of channels is encoded by eight genes, Kv10.1–10.2, Kv11.1–11.3, and Kv12.1–12.3, and many of these channels seem to be involved in cell cycle regulation (Pardo et al., 1998). In fact, the first discovery of their association with the cell cycle came from neuroblastoma. The whole-cell patch-clamp recording indicated that EAG current density varied widely from cell to cell (Meyer and Heinemann, 1998). However, if cells were synchronized in the G1 phase of the cell cycle, current density was the same in all cells. Furthermore, as cells moved through the cell cycle, current density changed with the largest density found in G1. This same phenomenon was later demonstrated for breast cancer cells as well (Ouadid-Ahidouch et al., 2001). Increasing the link between EAG cells and cancer means that EAG transcripts and/or currents can be found in a large number of cancers including neuroblastoma, breast, gastrointestinal, endometrial, and leukemic and colon cancers (Meyer et al., 1999; Smith et al., 2002; Suzuki and Takimoto, 2004; Ousingsawat et al., 2007). In the past 5 years, this list has grown to include gliomas (Patt et al., 2004). Specifically, both eag1 and erg1 transcripts have been discovered in patient biopsies of both low-and high-grade gliomas. Surprisingly, pharmacological inhibition of hERG currents appears to dramatically reduce secretion of vascular endothelial growth factor (VEGF) specifically as epidermal growth factor secretion was unaffected (Masi et al., 2005). Both growth factors are known to be secreted from gliomas and secretion of VEGF, in particular, has been of interest recently as a potential target to limit the growth of gliomas in vivo. Studies on EAG channels in gliomas are still in

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their infancy, but if the above results are substantiated, EAG channels may be the ideal Kþ channels to target therapeutically. Currently, specific inhibitors for these channels are unavailable, but given their intimate association with cancer, drug development to generate specific inhibitors is a worthy pursuit. 4.3 Cl
CHANNELS IN GLIOMAS A signature feature of nonmalignant glia is a very negative resting membrane potential (Kuffler, 1967), established primarily by the activity of the Kir4.1 inwardly rectifying Kþ channels. As discussed in the previous section, glioma cells do not express functional Kir channels and as a consequence have a relatively depolarized resting membrane potential of approximately
30 to
40 mV (Olsen and Sontheimer, 2004). We have already alluded to the fact that the absence of these channels is essential for cells to go through mitosis (Higashimori and Sontheimer, 2007). Surprisingly, however, the negative resting potential is at least in part established by a contribution of a resting Cl
conductance (Ransom et al., 2001). As illustrated in Fig. 4.3, when recorded with amphotericin patch clamp so as to not dialyze the cell, the application of the Cl
channel inhibitor NPPB caused the resting conductance to decrease markedly and input resistance to increase, while TEA had little effect. These data suggest that glioma cells indeed have Cl
channels that show significant open probability at rest and, secondly, that even at
40 mV, an outward-directed driving force exists for Cl
ions to leave the cells. The latter is particularly surprising as in many cell types, neurons being a prime example, Cl
conductance stabilizes the resting potential rather than depolarizes it, as is evident from the inhibitory action of GABA-gated Cl
channels (Ebihara et al., 1995). Using the fluorescence Cl
indicator SPQ, we have been able to demonstrate directly that intracellular Cl
in glioma cells is maintained around 80–100 mM through the activity of the NKCC1 transporter (unpublished). This value is at least twice as high as in astrocytes (Kimelberg, 1981) and approximately 10-fold higher than in most neurons (Ebihara et al., 1995). This fact will become important as we discuss the participation of Cl
channels in cell invasion, which requires an efflux of Cl
from these cells to promote cell shrinkage. Over the past 8 years, we have made a concerted effort to further identify and characterize the underlying Cl
channels in glioma cells. Cl
channels cloned to date fall into four molecular superfamilies (Valverde, 1999): the CFTR channels, Ca2þ-activated Cl
channels (Agnel et al., 1999), voltage-dependent anion-selective channels (VDACs) (Wunder and Colombini, 1991; Dermietzel et al., 1994; Dolder et al., 1999), and the ClC channels (Jentsch et al., 1999; Valverde, 1999). Of these, neither the CFTR nor Ca2þ-activated channels are present in gliomas but three members of the ClC family are consistently present in patient biopsies and isolated glioma cells (Olsen et al., 2003). These include ClC-2, ClC-3, and ClC-5. Biophysically, currents mediated by ClC-2 and ClC-3 channels could be identified. Unfortunately, specific inhibitors for Cl
channels are not available and hence most investigators rely on a combination of differential biophysical and pharmacological properties

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FIGURE 4.3 The resting chloride conductance of glioma cells is present in nondialyzed cells (amphotericin-perforated patch clamp). (a) Double y-axis plot of current (at
40 mV) and input resistance (Rin) as a function of time in an amphotericin-perforated patch-clamped cell. NPPB (165 mM), but not TEA (1 mM), blocked an inward current at typical resting potentials (
40 mV) and increased the input resistance. Inset shows current responses to a voltage step from
40 to
60 mV before and after NPPB application (used to calculate input resistance). The series resistance of this cell was 22 MW. (b) Summary of NPPB effects on input resistance in cells recorded from the perforated patch configuration. (c) Summary of NPPB effects on current at
40 mV in cells recorded from the perforated patch configuration. Data in (b and c) are displayed as mean  SE (Ransom et al., 2001).

to identify the underlying channels. We have complemented such approaches with the use of antisense oligonucleotides specific for each of the ClC channels of interest to more unequivocally identify currents as being mediated by ClC-2 and ClC-3, respectively. As illustrated in Fig. 4.4a and b, ClC-2 channels give rise to inwardly rectifying currents that are insensitive to NPPB but blocked by Cd2þof 200 mM. Currents are enhanced if voltage steps are preceded by a short hyperpolarization to
120 mV. ClC-3 channels, by contrast, give rise to currents that are outwardly rectifying and show time-dependent inactivation at positive potentials. Currents are inhibited by DIDS and NPPB but not Cd2þ. Currents with this identical biophysical signature are also inhibited by chlorotoxin, a 36 amino acid peptide isolated from Leiurus quinquestriatus and suggested to be a Cl
channel blocker (DeBin et al., 1993) (Fig. 4.4c). However, unlike NPPB, Cltx requires several minutes to achieve complete current inhibition (Ullrich et al., 1996). This discrepancy was recently explained by the

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FIGURE 4.4 Voltage-dependent inward Cl
currents in human glioma cells. (a) Inward currents were evoked with voltage steps from
140 to þ20 mV from a holding potential of
40 mV. Cells were hyperpolarized to
120 mV for a minimum of 20 s to increase the activation of inward current. Representative traces of inward Cl
current before (top) and after Cd2þ (200 mM) (middle) and the subtracted Cd2þ-sensitive current (bottom) are shown. (b) I–V plot of Cd2þ (200 mM)-sensitive Cl
currents evoked from the same voltage step protocol. Currents returned after washout of Cd2þ (Olsen et al., 2003) (c) Whole-cell currents were recorded before and after the application of 1 mM Cltx. A near complete inhibition of ClC-3 currents is achieved with Cltx. Of note, unlike the block by NPPB, which develops within seconds, complete block by Cltx requires 10–15 min, and the example illustrated is recorded 15 min after the application of Cltx (Weaver et al., 2007).

identification of the actual receptor for Cltx as a protein complex consisting of matrix metalloproteinase 2 (MMP-2), its receptor MT1-MMP, and the ClC-3 Cl
channel (Deshane et al., 2003; McFerrin and Sontheimer, 2005). Rather than blocking the channel directly, binding of Cltx to glioma cells causes the internalization of this molecular complex and hence the removal of functional ClC-3 channels from the plasma membrane into caveoli, a specialized vesicle associated with lipid rafts (McFerrin and Sontheimer, 2005). Although the underlying interactions and the signal that causes internalization of this complex remain elusive, the requirement for MMP-2 as target for Cltx has recently been confirmed in a related tumor model in vivo (Veiseh et al., 2007).

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In agreement with the hypothesized role of Cl
channels in the glioma cell shape changes that facilitate cell invasion, all inhibitors of ClC-3 have been shown to also effectively inhibit glioma cell invasion. The simplest way to mimic the spatial constraints encountered by invading glioma cells is through the use of the so-called transwell chambers, which are modified Boyden chambers in which cells are allowed to migrate through pores of 5 mm from one chamber to another toward a chemoattractant molecule. Replacement of Cl
in the medium with impermeant anions impeded the ability of cells to shrink sufficiently to transverse these barriers as did the inhibition of Cl
channels with NPPB, DIDS, or Cltx (Ransom et al., 2001). The latter also inhibited cell invasion in situ using a spheroid confrontation assay. Together with animal studies in which it was demonstrated that Cltx selectively homes in on the tumor cells (Soroceanu et al., 1999), these studies paved the way for a clinical use of Cltx in patients with malignant gliomas as discussed at the end of this review.

4.4 VOLTAGE-GATED SODIUM CHANNELS Voltage-gated sodium (Naþ) channels are responsible for the depolarizing upstroke of the action potential in neurons and muscle cells. This family of channels consists of nine alpha subunits (Nav1.1–1.9) that combine with two beta subunits to form a heterotrimer. Each of the nine alpha subunits is expressed in the brain and gives rise to voltage-dependent currents that rapidly inactivate. These channels are characterized by their sensitivity to tetrodotoxin (TTX), a toxin isolated from the puffer fish. Because these channels are the principal substrates for excitability, it came as a great surprise when Ritchie and colleagues demonstrated them also in glial cells (Ritchie, 1992), albeit at much lower density. Subsequent studies suggested that unlike neurons, astrocytic Naþ channels function to maintain the intracellular Naþ concentration at a level sufficient to fuel Naþ-Kþ-ATPase activity and thus energy production for the cell (Sontheimer et al., 1994). Most of the cancer researches are performed in cell lines that derive from patient biopsies and have often been cultured over hundreds of passages in the lab. Electrophysiological recordings from glioma cell lines indicated a lack of expression of voltage-gated Naþ channels. Since astrocytes often lose ion channel expression in passages in vitro, this prompted investigators to examine glioma cells in human brain slices and/or cells acutely dissociated from patient biopsies. With this approach, Patt et al. (1996) recorded whole-cell TTX-sensitive Naþ currents from oligodendroglioma and oligoastrocytoma cells (Fig. 4.5a). Interestingly, oligodendrocytes, the presumed precursor to the tumors examined in this study, do not normally express Naþ channels leading one to speculate that the expression of these channels may be integral to the malignant transformation and propagation of these tumors. Bordey and Sontheimer (1998a) independently reported similar findings from astrocytomas. Specifically, the whole-cell patch recordings of low grade astrocytomas identified Naþ currents identical to those found in oligodendrogliomas. Possibly the most intriguing discovery of both of

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FIGURE 4.5 Astrocytoma cells express tetrodotoxin (TTX)-sensitive, voltage-dependent Naþ currents. (a) Whole-cell inward currents in response to 8 ms step depolarizations starting at
70 mV incremented by 10 mV from a holding potential of
110 mV. Currents before exposure to TTX (left) and after a 3-min exposure to 100 nM TTX (right). (b) Current–clamp traces from a glioma cell of a tumor biopsy after a short depolarizing current pulse. Spike amplitude was 52 mV, threshold was approximately
33 mV, rise time (beginning of pulse to peak value) was 15.1 ms, duration of spike from threshold was 11.3 ms, and corresponding ratio for the same recorded cell was 1.1 (Adapted from Bordey and Sontheimer, 1998a).

these studies is that upon injection of current, glioma cells are capable of generating spikes (Fig. 4.5b) akin to neuronal action potentials. However, the half-maximal voltage for inactivation of these channels in glioma cells is
47 mV, making it highly unlikely that these cells could ever generate an action potential in vivo since the typical resting membrane potential of a glioma cell is 
40 mV. What makes this finding quite interesting is the fact that glioma cells express Naþ channels at a sufficient density to generate action potentials and hence express a much larger number of Naþ channels than that found in the nonmalignant glia. The reasons for this are entirely unclear. However, given the proliferative capacity of glioma cells, it has been speculated that these channels may serve a similar function in gliomas as in astrocytes, that is, supporting the activity of the cells Naþ/Kþ-ATPase. Thus,

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because glioma cells are metabolically quite active, a greater Naþ influx may be required to sustain their Naþ-Kþ-ATPase. Attempts have been made, albeit unsuccessful, to identify the subunit responsible for the Naþ currents in glioma cells. Glioma cell lines and biopsies, alike, seem to variably express each of the nine Naþ channel alpha subunits (Schrey et al., 2002). Moreover, functional studies are difficult since expression of these channels is lost in culture (Patt et al., 1996). A recent study was able to recover Naþ channel expression in glioma cell lines through a combination of removal of the fetal calf serum that is normally present in culture media and addition of nerve growth factor (NGF) (Kraft et al., 2001). In the future, this approach may not only make possible to assess the function of voltage-gated Naþ channels in gliomas, but may also allow to study pathways responsible for their regulation. However, regardless of the function of Naþ channels in gliomas, they make poor pharmacological targets due to their ubiquitous expression in the brain and their importance in all aspects of nervous system function.

4.5 VOLTAGE-GATED CALCIUM CHANNELS Arguably, the most ubiquitous ion channel expressed and found in almost all cell types and animal species are the voltage-gated calcium channels (VGCCs). These are comprised of a pore-forming a1 subunit along with three or four additional modulatory subunits. Five members of the VGCC family are encoded by 10 genes and thus there are 10 different a1 subunits: the L-type channels are encoded by Cav1.1–1.4, the N-, P/Q-, and R-type channels are encoded by Cav2.1–2.3, respectively, and the T-type channels are encoded by Cav3.1–3.3. The L-, N-, P/Q- and Rtype channels give rise to currents that are activated by large depolarizations and hence are collectively called the high-voltage activated (HVA) calcium channels. The L-type VGCCs can be blocked by a family of compounds termed dihydropyridines that include verapamil, nifedipine, and nicardipine. The remaining HVA channels each exhibit unique pharmacology; however, all HVA channels are sensitive to the heavy metal Cd2þ. In contrast, the T-type calcium channels are activated by very small depolarizations and give rise to small, transient currents. These channels are typically referred to as low-voltage activated (LVA) channels. In addition to being characterized by unique voltage dependence, LVA channels are not sensitive to typical VGCC antagonists and are instead inhibited by the heavy metal Ni2þ and the organic compound mibefradil. Much like Naþ channels, VGCCs can be found in all excitable cells where they have well-defined roles in action potential generation and neurotransmitter release. Indeed, therapeutically targeting VGCCs has proven extremely successful in the treatment of cardiac disorders such as hypertension and arrhythmia. In nonexcitable cells, these channels have been shown to be involved in both cell proliferation and cell migration. In the mid-1980s, it was discovered that coapplication of the L-type VGCC inhibitors verapamil, nifedipine, or nicardipine with various chemotherapeutic agents actually enhanced the cytotoxic effect of these drugs and in

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some cases overcame the adaptive resistance that gliomas can develop against chemotherapy drugs (Kaba et al., 1985; Merry et al., 1986; Bowles et al., 1990; Kondo et al., 1995). Interestingly, calcium imaging in the presence or absence of one of the L-type VGCC inhibitors, verapamil, indicated a lack of verapamil-sensitive calcium fluxes in glioma cells (Huet and Robert, 1988), leading researchers to conclude that calcium channel blockers exert their chemotherapeutic-enhancing effects by a mechanism independent of its inhibition of VGCCs. Subsequent research suggested that verapamil actually downregulated the expression of P-glycoprotein, a protein thought to be responsible for the multidrug-resistant phenotype of gliomas (Kiwit et al., 1994; Abe et al., 1995), although recent evidence has questioned whether this is actually the case (Huet et al., 1992; Rieger et al., 2000). Aside from their direct effect on tumor cells, the L-type VGCC inhibitors may act on other cell types to enhance drug delivery to the tumor site. Specifically, two studies from the late 1990s indicate that intravenous administration of nimodipine may increase blood–tumor barrier (BTB) selectively, leaving the BTB unaffected (Matsukado et al., 1994) and allowing higher concentrations of chemotherapeutic agents to reach the tumor specifically (Zenke et al., 1996). Regardless of the mechanism by which verapamil, minodipine, and nicardipine act to enhance cytotoxity of common anticancer drugs and despite the seeming lack of L-type VGCC expression in gliomas, the use of their inhibitors may offer a new twist on conventional therapies to enhance their outcome. Indeed, several animal studies indicate that combining the common chemotherapy treatment with the L-type VGCC inhibitors might be a successful approach to combating gliomas. Analysis of mRNA transcripts has revealed that gliomas express Cav3.1, a T-type VGCC (Latour et al., 2004). Furthermore, this channel appears to be a unique splice variant in that when exogenously expressed, it appears to have slower inactivation kinetics compared to the known Cav3.1 channels. Emerging research is indicating that of all the VGCCs, the T-type channel seems to be expressed in several cancers (Panner and Wurster, 2006). Indeed, a recent study has indicated that the T-type channels may be involved in the glioma proliferation as pharmacologic inhibition with mibefradil, the most widely used T-type channel inhibitor, and knockdown of the channel with antisense oligonucleotides reduces glioma proliferation by 50 % (Panner et al., 2005). The involvement of T-type calcium channels in the proliferation of cancer cells may not be surprising. Unlike the other VGCCs, the T-type channels activate in response to much smaller depolarizations that are reasonably achieved in nonexcitable cells. Furthermore, these channels have been reported to be regulated by calmodulin in addition to voltage (Barrett et al., 2000). Calmodulin is a common signaling molecule activated by growth factors known to be constitutively active in gliomas. The potential modulation of these ion channels by growth factors positions them as the primary calcium entry pathway in proliferative signaling cascades. Evidence for this already exists in gliomas. Specifically, platelet-derived growth factor–BB (PDGF–BB) induces a sustained Ca2þ increase in glioma cell lines that can be inhibited by the T-type channel inhibitor NDGA (Saqr et al., 1999). Increases in [Ca2þ]i are absolutely required for cell cycling. Despite a lack of electrophysiological evidence for the

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functional T-type VGCCs, molecular biology and pharmacology indicate that these channels may be intricately involved in these proliferative calcium signals and, therefore, might be potential therapeutic targets. Unfortunately, specific pharmacological inhibitors of the T-type channels do not currently exist. However, the implication of their involvement in multiple cancers as well as various other disorders such as chronic pain and epilepsy is driving a concerted effort to develop specific inhibitors as future therapeutic tools.

FIGURE 4.6 (a) Model representing glioma cell shape and volume-adaptive changes that occur during invasion in spatially restricted conditions. These changes, accompanied by water loss and cytoskeletal rearrangements, are mediated by ion fluxes through Cl
and Kþ ion channels and other ion transport mechanisms. (b) Immunocytochemistry demonstrates a colocalization of both ClC3 channel protein and BK channel protein with the cholera toxin-B subunit as evidenced by yellow staining in the merged image. This subunit specifically binds to GM1 ganglioside, a well-characterized component of lipid rafts. (c) Biochemical separation of lipid raft proteins indicates that both ClC3 and BK channels can be found in the second detergent insoluble fraction of one such separation. This fraction contains common lipid raft proteins including Caveolin-1, a marker of a subset of lipid rafts. (Weaver et al., 2007). (See the color version of this figure in Color Plate section.)

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4.6 USE OF CHANNEL INHIBITORS FOR THE TREATMENT OF GLIOMAS It becomes quite evident from the above studies that several ion channels are prominently expressed in glioma cells and appear to be necessary for distinct aspects of glioma cell biology. BK channels appear to confer an advantage to cells to survive in conditions of growth factor starvation. The downregulation of Kir4.1 appears necessary for cells to maintain cycling. Similarly, it is clear that inhibition of certain channels, gBK channels or ClC-3 Cl
channels, impedes the ability of cells to invade in vitro. While the mechanistic interdependence of channel function and the underlying biology are not always obvious, a good mechanistic model can be presented to explain how Kþ and Cl
channels collaborate to support shape changes of invading cells. As illustrated in Fig. 4.6a, fluid dynamics is the mechanistic underpinning in that a coordinated flux of KCl and water is the requirement to attain a cell volume conducive to cell invasion. Not surprisingly therefore, all channels involved in fluid secretion colocalize to the invading aspects of cells where they are clustered in specialized lipid raft domains (Fig. 4.6b and c). Kþ and Cl
channels may serve a similar role in other inexcitable cells including other cancer cells. This complex of channels, localized to lipid rafts, may provide an opportune therapeutic target. Unfortunately, blockers for BK channels lack glioma selectivity and useful aquaporin blockers do not exist. Fortunately, the putative Cl
channel blocker Cltx causes functional inhibition of Cl
currents by disrupting the membrane association of ClC-3 channels. Since Cltx binds almost exclusively to the cancer cells in the brain (Soroceanu et al., 1998) and lacks specific binding to normal human organ tissues, its clinical use was proposed for targeting experiments in animal models. Clinical grade Cltx has now been manufactured and introduced into a phase I clinical study that reported its findings recently (Mamelak et al., 2006). As illustrated in Fig. 4.7, Cltx shows remarkable glioma specificity in patients where the molecule is retained at the

FIGURE 4.7 Axial view of T1-Wc, (a) coregistered (b) and SPECT (c) (day 8) images. Longterm retention of 131I-TM-601 radioactivity at tumor site was observed in all patient scans (Hockaday et al., 2005). (See the color version of this figure in Color Plate section.)

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tumor site for up to 8 days. This retention indicates that Cltx is endocytosed and trapped inside the glioma cell in vivo analogous to our previous culture experiments. Importantly, the phase I clinical trial established drug safety and paved the way for a multicenter phase II clinical trial that is currently enrolling patients with malignant glioma. ACKNOWLEDGMENTS This work was supported by NIH grants RO1-NS031234, RO1-NS036692, and P50CA97247.

REFERENCES Abe T, Koike K, Ohga T, Kubo T, Wada M, Kohno K, Mori T, Hidaka K, Kuwano M, 1995. Chemosensitisation of spontaneous multidrug resistance by a 1,4-dihydropyridine analogue and verapamil in human glioma cell lines overexpressing mrp or mdr1. Br J Cancer 72: 418–423. Agnel M, Vermat T, Culouscou JM, 1999. Identification of three novel members of the calcium-dependent chloride channel (CaCC) family predominantly expressed in the digestive tract and trachea. FEBS Lett 455: 295–301. Barrett PQ, Lu HK, Colbran R, Czernik A, Pancrazio JJ, 2000. Stimulation of unitary T-type Ca2þ channel currents by calmodulin-dependent protein kinase II. Am J Physiol Cell Physiol 279: C1694–C1703. Basrai D, Kraft R, Bollensdorff C, Liebmann L, Benndorf K, Patt S, 2002. BK channel blockers inhibit potassium-induced proliferation of human astrocytoma cells. Neuroreport 13: 403–407. Belien AT, Paganetti PA, Schwab ME, 1999. Membrane-type 1 matrix metalloprotease (MT1MMP) enables invasive migration of glioma cells in central nervous system white matter. J Cell Biol 144: 373–384. Bordey A, Sontheimer H, 1997. Postnatal development of ionic currents in rat hippocampal astrocytes in situ. J Neurophysiol 78: 461–477. Bordey A, Sontheimer H, 1998a. Electrophysiological properties of human astrocytic tumor cells in situ: enigma of spiking glial cells. J Neurophysiol 79: 2782–2793. Bordey A, Sontheimer H, 1998b. Passive glial cells, fact or artifact? J Memr Biol 166: 213–222. Bordey A, Sontheimer H, 2000. Ion channel expression by astrocytes in situ: comparison of different CNS regions. Glia 30(1): 27–38. Bordey A, Sontheimer H, Trouslard J, 2000. Muscarinic activation of BK channels induces membrane oscillations in glioma cells and leads to inhibition of cell migration. J Memr Biol 176: 31–40. Bordey A, Lyons SA, Hablitz JJ, Sontheimer H, 2001. Electrophysiological characteristics of reactive astrocytes in experimental cortical dysplasia. J Neurophysiol. Bowles AP Jr, Pantazis CG, Wansley W, 1990. Use of verapamil to enhance the antiproliferative activity of BCNU in human glioma cells: an in vitro and in vivo study. J Neurosurg 73: 248–253.

REFERENCES

121

Camacho J, 2006. Ether a go-go potassium channels and cancer. Cancer Lett 233: 1–9. Chicoine MR, Silbergeld DL, 1997. Mitogens as motogens. J Neurooncol 35: 249–257. Chvatal A, Pastor A, Mauch M, Sykova E, Kettenmann H, 1995. Distinct populations of identified glial cells in the developing rat spinal cord slice: ion channel properties and cell morphology. Eur J Neurosci 7: 129–142. Collins VP, 1995. Gene amplification in human gliomas. Glia 15: 289–296. DeBin JA, Maggio JE, Strichartz GR, 1993. Purification and characterization of chlorotoxin, a chloride channel ligand from the venom of the scorpion. Am J Physiol 264: C361–C369. Demuth T, Berens ME, 2004. Molecular mechanisms of glioma cell migration and invasion. J Neurooncol 70: 217–228. Dermietzel R, Hwang T-K, Buettner R, Hofer A, Dotzler E, Kremer M, Deutzmann R, Thinnes FP, Fishman GI, Spray DC, Siemen D, 1994. Cloning and in situ localization of a brain-derived porin that constitutes a large-conductance anion channel in astrocytic plasma membranes. Proc Natl Acad Sci USA 91: 499–503. Deryugina EI, Bourdon MA, Luo GX, Reisfeld RA, Strongin A, 1997. Matrix metalloproteinase-2 activation modulates glioma cell migration. J Cell Sci 110: 2473–2482. Deshane J, Garner CC, Sontheimer H, 2003. Chlorotoxin inhibits glioma cell invasion via matrix metalloproteinase-2. J Biol Chem 278: 4135–4144. Di X, Goforth PB, Bullock R, Ellis E, Satin L, 2000. Mechanical injury alters volume activated ion channels in cortical astrocytes. Acta Neurochir Suppl 76: 379–383. Dolder M, Zeth K, Tittmann P, Gross H, Welte W, Wallimann T, 1999. Crystallization of the human, mitochondrial voltage-dependent anion-selective channel in the presence of phospholipids. J Struct Biol 127: 64–71. Ebihara S, Shirato K, Harata N, Akaike N, 1995. Gramicidin-perforated patch recording: GABA response in mammalian neurones with intact intracellular chloride. J Physiol 484 (Part 1): 77–86. Farin A, Suzuki SO, WeikerM, Goldman JE, BruceJN, Canoll P, 2006. Transplantedgliomacells migrate and proliferate on host brain vasculature: a dynamic analysis. Glia 53: 799–808. Giese A, Rief MD, Loo MA, Berens ME, 1994. Determinants of human astrocytoma migration. Cancer Res 54: 3897–3904. Giese A, Loo MA, Rief MD, Tran N, Berens ME, 1995. Substrates for astrocytoma invasion. Neurosurgery 37: 294–301. Giese A, Laube B, Zapf S, Mangold U, Westphal M, 1998. Glioma cell adhesion and migration on human brain sections. Anticancer Res 18: 2435–2447. Gladson CL, 1999. The extracellular matrix of gliomas: modulation of cell function. J Neuropathol Exp Neurol 58: 1029–1040. Higashi K, Fujita A, Inanobe A, Tanemoto M, Doi K, Kubo T, Kurachi Y, 2001. An inwardly rectifying Kþ channel, Kir4.1, expressed in astrocytes surrounds synapses and blood vessels in brain. Am J Physiol Cell Physiol 281: C922–C931. Higashimori H, Sontheimer H, 2007. Role of Kir4.1 Channels in growth control of glia. Glia 55:1668–1679. Hockaday DC, Shen S, Fiveash J, Raubitschek A, Colcher D, Liu A, Alvarez V, Mamelak AN, 2005. Imaging glioma extent with 131I-TM601. J Nucl Med 46(4): 580–586. Huet S, Robert J, 1988. The reversal of doxorubicin resistance by verapamil is not due to an effect on calcium channels. Int J Cancer 41: 283–286.

122

THE ROLE OF ION CHANNELS IN THE ETIOLOGY AND DEVELOPMENT OF GLIOMAS

Huet S, Schott B, Robert J, 1992. P-glycoprotein overexpression cannot explain the complete doxorubicin-resistance phenotype in rat glioblastoma cell lines. Br J Cancer 65: 538–544. Jentsch TJ, Friedrich T, Schriever A, Yamada H, 1999. The CLC chloride channel family. Pfl€ ugers Arch 437: 783–795. Kaba K, Tani E, Morimura T, Matsumoto T, 1985. Potentiation of vincristine effect in human and murine gliomas by calcium channel blockers or calmodulin inhibitors. J Neurosurg 63: 905–911. Kimelberg HK, 1981. Active accumulation and exchange transport of chloride in astroglial cells in culture. Biochim Biophys Acta 646: 179–184. Kiwit JC, Hertel A, Matuschek AE, 1994. Reversal of chemoresistance in malignant gliomas by calcium antagonists: correlation with the expression of multidrug-resistant P-glycoprotein. J Neurosurg 81: 587–594. Kleihues P, Ohgaki H, Aguzzi A, 1995. Gliomas. In: Kettenmann H, Ransom BR (Eds), Neuroglia, Oxford University Press, pp. 1044–1063. Kofuji P, Biedermann B, Siddharthan V, Raap M, Iandiev I, Milenkovic I, Thomzig A, Veh RW, Bringmann A, Reichenbach A, 2002. Kir potassium channel subunit expression in retinal glial cells: implications for spatial potassium buffering. Glia 39: 292–303. Kondo S, Yin D, Morimura T, Takeuchi J, 1995. Combination therapy with cisplatin and nifedipine inducing apoptosis in multidrug-resistant human glioblastoma cells. J Neurosurg 82: 469–474. Kraft R, Basrai D, Benndorf K, Patt S, 2001. Serum deprivation and NGF induce and modulate voltage-gated Naþ currents in human astrocytoma cell lines. Glia 34: 59–67. Kraft R, Krause P, Jung S, Basrai D, Liebmann L, Bolz J, Patt S, 2003. BK channel openers inhibit migration of human glioma cells. Pflugers Arch 446: 248–255. Kucheryavykh YV, Kucheryavykh LY, Nichols CG, Maldonado HM, Baksi K, Reichenbach A, Skatchkov SN, Eaton MJ, 2007. Downregulation of Kir4.1 inward rectifying potassium channel subunits by RNAi impairs potassium transfer and glutamate uptake by cultured cortical astrocytes. Glia 55: 274–281. Kuffler SW, 1967. Neuroglial cells: physiological properties and a potassium mediated effect of neuronal activity on the glial membrane potential. Proc R Soc Lond B:Biol Sci 168: 1–21. Kunzelmann K, 2005. Ion channels and cancer. J Membr Biol 205: 159–173. Latour I, Louw DF, Beedle AM, Hamid J, Sutherland GR, Zamponi GW, 2004. Expression of T-type calcium channel splice variants in human glioma. Glia 48: 112–119. Li L, Head V, Timpe LC, 2001. Identification of an inward rectifier potassium channel gene expressed in mouse cortical astrocytes. Glia 33: 57–71. Liu X, Chang Y, Reinhart PH, Sontheimer H, Chang Y, 2002. Cloning and characterization of glioma BK, a novel BK channel isoform highly expressed in human glioma cells. J Neurosci 22: 1840–1849. MacFarlane SN, Sontheimer H, 1997. Electrophysiological changes that accompany reactive gliosis in vitro. J Neurosci 17: 7316–7329. MacFarlane SN, Sontheimer H, 2000a. Changes in ion channel expression accompany cell cycle progression of spinal cord astrocytes. Glia 30(1): 39–48. MacFarlane SN, Sontheimer H, 2000b. Modulation of kv1.5 currents by Src tyrosine phosphorylation: potential role in the differentiation of astrocytes. J Neurosci 20(14): 5245–5253.

REFERENCES

123

Maidment SL, Merzak A, Koochekpour S, Rooprai HK, Rucklidge GJ, Pilkington GJ, 1996. The effect of exogenous gangliosides on matrix metalloproteinase secretion by human glioma cells in vitro. Eur J Cancer 32A, 868–871. Maidment SL, Rucklidge GJ, Rooprai HK, Pilkington GJ, 1997. An inverse correlation between expression of NCAM-A and the matrix-metalloproteinases gelatinase-A and gelatinase-B in human glioma cells in vitro. Cancer Lett 116: 71–77. Mamelak AN, Rosenfeld S, Bucholz R, Raubitschek A, Nabors LB, Fiveash JB, Shen S, Khazaeli MB, Colcher D, Liu A, Osman M, Guthrie B, Schade-Bijur S, Hablitz DM, Alvarez VL, Gonda MA, 2006. Phase I single-dose study of intracavitary-administered iodine-131-TM-601 in adults with recurrent high-grade glioma. J Clin Oncol 24: 3644–3650. Manning TJ Jr, Parker JC, Sontheimer H, 2000. Role of lysophosphatidic acid and rho in glioma cell motility. Cell Motil Cytoskeleton 45(3): 185–199. Masi A, Becchetti A, Restano-Cassulini R, Polvani S, Hofmann G, Buccoliero AM, Paglierani M, Pollo B, Taddei GL, Gallina P, Di Lorenzo N, Franceschetti S, Wanke E, Arcangeli A, 2005. hERG1 channels are overexpressed in glioblastoma multiforme and modulate VEGF secretion in glioblastoma cell lines. Br J Cancer 93: 781–792. Matsukado K, Nomura T, Ikezaki K, Fukui M, 1994. Selective increase in blood–tumor barrier permeability by calcium antagonists in transplanted rat brain tumors. Acta Neurochir Suppl (Wien) 60: 403–405. McFerrin MB, Sontheimer H, 2005. A role for ion channels in glioma cell invasion. Neuron Glia Biol 2: 39–49. Merry S, Fetherston CA, Kaye SB, Freshney RI, Plumb JA, 1986. Resistance of human glioma to adriamycin in vitro: the role of membrane transport and its circumvention with verapamil. Br J Cancer 53: 129–135. Merzak A, McCrea S, Koocheckpour S, Pilkington GJ, 1994. Control of human glioma cell growth, migration and invasion in vitro by transforming growth factor b1. Br J Cancer 70: 199–203. Meyer R, Heinemann SH, 1998. Characterization of an eag-like potassium channel in human neuroblastoma cells. J Physiol (Lond) 508: 49–56. Meyer R, Sch€onherr R, Gavrilova-Ruch O, Wohlrab W, Heinemann SH, 1999. Identification of ether a go-go and calcium-activated potassium channels in human melanoma cells. J Memr Biol 171: 107–115. Nakada M, Okada Y, Yamashita J, 2003. The role of matrix metalloproteinases in glioma invasion. Front Biosci 8: E261–E269. Neusch C, Papadopoulos N, Muller M, Maletzki I, Winter SM, Hirrlinger J, Handschuh M, Bahr M, Richter DW, Kirchhoff F, Hulsmann S, 2006. Lack of the Kir4.1 channel subunit abolishes Kþ buffering properties of astrocytes in the ventral respiratory group: impact on extracellular Kþ regulation. J Neurophysiol 95: 1843–1852. Ohgaki H, Kleihues P, 2005. Population-based studies on incidence, survival rates, and genetic alterations in astrocytic and oligodendroglial gliomas. J Neuropathol Exp Neurol 64: 479–489. Ohnishi T, Arita N, Hayakawa T, Kawahara K, Kato K, Kakinuma A, 1993. Purification of motility factor (gmf) from human malignant glioma cells and its biological significance in tumor invasion. Biochem Biophys Res Commun 193: 518–525. Olsen ML, Sontheimer H, 2004. Mislocalization of Kir channels in malignant glia. Glia 46: 63–73.

124

THE ROLE OF ION CHANNELS IN THE ETIOLOGY AND DEVELOPMENT OF GLIOMAS

Olsen ML, Schade S, Lyons SA, Amarillo MD, Sontheimer H, 2003. Expresssion of voltagegated chloride channels in human glioma cells. J Neurosci 23: 5572–5582. Olsen ML, Weaver AK, Ritch PS, Sontheimer H, 2005. Modulation of glioma BK channels via erbB2. J Neurosci Res 81: 179–189. Olsen ML, Higashimori H, Campbell SL, Hablitz JJ, Sontheimer H, 2006. Functional expression of K(ir)4.1 channels in spinal cord astrocytes. Glia 53: 516–528. Olsen ML, Campbell SL, Sontheimer H, 2007. Differential distribution of Kir4.1 in spinal cord astrocytes suggests regional differences in kþ homeostasis. J Neurophysiol 98: 786–793. Ouadid-Ahidouch H, Le BX, Roudbaraki M, Toillon RA, Delcourt P, Prevarskaya N, 2001. Changes in the Kþ current-density of MCF-7 cells during progression through the cell cycle: possible involvement of a h-ether-a-go-go Kþ channel. Receptors Channels 7: 345–356. Ousingsawat J, Spitzner M, Puntheeranurak S, Terracciano L, Tornillo L, Bubendorf L, Kunzelmann K, Schreiber R, 2007. Expression of voltage-gated potassium channels in human and mouse colonic carcinoma. Clin Cancer Res 13: 824–831. Panner A, Wurster RD, 2006. T-type calcium channels and tumor proliferation. Cell Calc 40: 253–259. Panner A, Cribbs LL, Zainelli GM, Origitano TC, Singh S, Wurster RD, 2005. Variation of T-type calcium channel protein expression affects cell division of cultured tumor cells. Cell Calc 37: 105–119. Pardo LA, Bruggemann A, Camacho J, Stuhmer W, 1998. Cell cycle-related changes in the conducting properties of r-eag Kþ channels. J Cell Biol 143: 767–775. Parkerson KA, Sontheimer H, 2004. Biophysical and pharmacological characterization of hypotonically activated chloride currents in cortical astrocytes. Glia 46: 419–436. Patt S, Labrakakis C, Bernstein M, Weydt P, Cervos-Navarro J, Nisch G, Kettenmann H, 1996. Neuron-like physiological properties of cells from human oligodendroglial tumors. Neuroscience 71: 601–611. Patt S, Preussat K, Beetz C, Kraft R, Schrey M, Kalff R, Schonherr K, Heinemann SH, 2004. Expression of ether a go-go potassium channels in human gliomas. Neurosci Lett 368: 249–253. Ransom CB, Sontheimer H, 1995. Biophysical and pharmacological characterization of inwardly rectifying Kþ currents in rat spinal cord astrocytes. J Neurophysiol 73: 333–345. Ransom CB, Sontheimer H, 2001. BK channels in human glioma cells. J Neurophysiol 85: 790– 803. Ransom CB, O’Neal JT, Sontheimer H, 2001. Volume-activated chloride currents contribute to the resting conductance and invasive migration of human glioma cells. J Neurosci 21: 7674– 7683. Ransom CB, Liu X, Sontheimer H, 2002. BK channels in human glioma cells have enhanced calcium sensitivity. Glia 38: 281–291. Rieger L, Rieger J, Winter S, Streffer J, Esser P, Dichgans J, Meyermann R, Weller M, 2000. Evidence for a constitutive, verapamil-sensitive, non-P-glycoprotein multidrug resistance phenotype in malignant glioma that is unaltered by radiochemotherapy in vivo. Acta Neuropathol (Berl) 99: 555–562. Ritch PA, Carroll SL, Sontheimer H, 2003. Neuregulin-1 enhances motility and migration of human astrocytic glioma cells. J Biol Chem 278: 20971–20978. Ritchie JM, 1992. Voltage-gated ion channels in Schwann cells and glia. TINS 15: 345–351. Saqr HE, Guan Z, Yates AJ, Stokes BT, 1999. Mechanisms through which PDGF alters intracellular calcium levels in U-1242 MG human glioma cells. Neurochem Int 35: 411–422.

REFERENCES

125

Schrey M, Codina C, Kraft R, Beetz C, Kalff R, Wolfl S, Patt S, 2002. Molecular characterization of voltage-gated sodium channels in human gliomas. Neuroreport 13: 2493–2498. Smith GA, Tsui HW, Newell EW, Jiang X, Zhu XP, Tsui FW, Schlichter LC, 2002. Functional up-regulation of HERG Kþ channels in neoplastic hematopoietic cells. J Biol Chem 277: 18528–18534. Sontheimer H, Waxman SG, 1993. Expression of voltage-activated ion channels by astrocytes and oligodendrocytes in the hippocampal slice. J Neurophysiol 70: 1863–1873. Sontheimer H, Minturn JE, Black JA, Ransom BR, Waxman SG, 1991. Two types of Naþcurrents in cultured rat optic nerve astrocytes: changes with time in culture and with age of culture derivation. J Neurosci Res 30: 275–287. Sontheimer H, Fernandez-Marques E, Ullrich N, Pappas CA, Waxman SG, 1994. Astrocyte Naþ channels are required for maintenance of Naþ/Kþ-ATPase activity. J Neurosci 14: 2464–2475. Sontheimer H, Black JA, Waxman SG, 1996. Voltage-gated Naþ channels in glia: properties and possible functions. Trends Neurosci 19: 325–331. Soroceanu L, Gillespie Y, Khazaeli MB, Sontheimer H, 1998. Use of chlorotoxin for targeting of primary brain tumors. Cancer Res 58: 4871–4879. Soroceanu L, Manning TJ Jr, Sontheimer H, 1999. Modulation of glioma cell migration and invasion using Cl
and Kþ ion channel blockers. J Neurosci 19: 5942–5954. Suzuki T, Takimoto K, 2004. Selective expression of HERG and Kv2 channels influences proliferation of uterine cancer cells. Int J Oncol 25: 153–159. Tonn JC, Goldbrunner R, 2003. Mechanisms of glioma cell invasion. Acta Neurochir Suppl 88: 32. Treasurywala S, Berens ME, 1998. Migration arrest in glioma cells is dependent on the av integrin subunit. Glia 24: 236–243. Uhm JH, Gladson CL, Rao JS, 1999. The role of integrins in the malignant phenotype of gliomas. Front Biosci 4: 59. Ullrich N, Gillespie GY, Sontheimer H, 1996. Human astrocytoma cells express a unique chloride current. Neuroreport 7: 1020–1024. Ullrich N, Bordey A, Gillespie GY, Sontheimer H, 1998. Expression of voltage-activated chloride currents in acute slices of human gliomas. Neuroscience 83: 1161–1173. Valverde MA, 1999. Cic channels: leaving the dark ages on the verge of a new millennium. Curr Opin Cell Biol 11: 509–516. [review, 82 refs]. Veiseh M, Gabikian P, Bahrami SB, Veiseh O, Zhang M, Hackman RC, Ravanpay AC, Stroud MR, Kusuma Y, Hansen SJ, Kwok D, Munoz NM, Sze RW, Grady WM, Greenberg NM, Ellenbogen RG, Olson JM, 2007. Tumor paint: a chlorotoxin:Cy5.5 bioconjugate for intraoperative visualization of cancer foci. Cancer Res 67: 6882–6888. Visted T, Enger PO, Lund-Johansen M, Bjerkvig R, 2003. Mechanisms of tumor cell invasion and angiogenesis in the central nervous system. Front Biosci 8: 32. Weaver AK, Liu X, Sontheimer H, 2004. Role for calcium-activated potassium channels (BK) in growth control of human malignant glioma cells. J Neurosci Res 78: 224–234. Weaver AK, Bomben VC, Sontheimer H, 2006. Expression and function of calcium-activated potassium channels in human glioma cells. Glia 54: 223–233. Weaver AK, Olsen ML, McFerrin MB, Sontheimer H, 2007. BK channels are linked to IP3receptors via lipid rafts: A novel mechanism for coupling [Ca2þ]i to ion channel activation. J Biol Chem 282: 31558–31568.

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Wonderlin WF, Strobl JS, 1996. Potassium channels, proliferation and G1 progression 385. J Membr Biol 154: 91–107. Wunder UR, Colombini M, 1991. Patch clamping VDAC in liposomes containing whole mitochondrial membranes. J Memr Biol 123: 83–91. Zamecnik J, 2005. The extracellular space and matrix of gliomas. Acta Neuropathol (Berl) 110: 435–442. Zenke K, Nakagawa K, Kumon Y, Ohta S, Hatakeyama T, Sakaki S, 1996. A strategy for selective anti-cancer drug concentration increase in rat glioma tissue with Ca2þ-channel blocker co-administration: calcium kinetics in intra-glioma arteriolar smooth muscle cells. J Neurooncol 30: 25–36.

5 SHAKER FAMILY KV1 VOLTAGEGATED POTASSIUM CHANNELS IN MAMMALIAN BRAIN NEURONS HELENE VACHER1 AND JAMES S. TRIMMER1,2 1

Department of Neurobiology, Physiology, and Behavior, College of Biological Sciences, University of California, Davis, CA 95616, USA 2

Department of Physiology and Membrane Biology, School of Medicine, University of California, Davis, CA 95616, USA

5.1 INTRODUCTION TO SHAKER FAMILY VOLTAGE-GATED POTASSIUM OR KV1 CHANNELS 5.1.1

KV1 Channels and Neuronal Excitability

Electrical excitability is a fundamental property of neurons. Diversity in intrinsic neuronal excitability and function is generated by the variable expression, subcellular localization, and activity of a complex repertoire of neuronal ion channels (Llinas, 1988). Dynamic regulation of intrinsic excitability can further alter the behavior of neurons and confer plasticity to neuronal signaling (Mohapatra et al., 2007). Moreover, aberrant expression, localization, and function of ion channels can result in channel-based pathophysiologies or channelopathies (Cannon, 2006). Thus, an understanding of how neurons regulate the expression and localization of ion channels is critical to understanding the complexity of normal neuronal function, its dynamic modulation to achieve plasticity, and defects that lead to neuronal dysfunction. Among the large complement of ion channels expressed in mammalian neurons are an especially wide variety of voltage-dependent potassium (Kv) channels. Among Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

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these, members of the Kv1 subfamily of Kv channels regulate diverse neuronal electrical properties, including action potential amplitude and duration, the frequency of cell firing, the kinetics and amount of neurotransmitter release, and the cells resting membrane potential (Dodson and Forsythe, 2004). Kv1 channels are supramolecular protein complexes composed of four pore-forming and voltage-sensing principal, or a, subunits plus up to four associated auxiliary or Kvb subunits (Pongs, 1999). In heterologous cells, these Kv1a and Kvb subunits can heteromultimerize to yield biophysically and pharmacologically distinct a4b4 channel complexes (Pongs, 1999). Kv1 channels can also associate with various scaffolding proteins and enzymes, which can impact channel localization, turnover, and function (Schulte et al., 2006). As such, the functional characteristics, abundance, and subcellular localization of Kv channel complexes are determined by diverse protein–protein interactions, both between constituent Kv1a and Kvb subunits, and between these subunits and a wide variety of interacting proteins (Levitan, 2006). 5.1.2

Expression of Kv1 Channel Subunits

Expression of Kv1 channel genes is highly regulated, with specific promoter elements acting in concert with transcriptional machinery to achieve precise temporal and spatial cellular patterns of expression (Mandel and McKinnon, 1993). Although evidence for dynamic regulation of ion channel translation has not been provided, it is clear that multiple posttranslational processes, including protein–protein interactions mediating subunit assembly, folding, and ultimately export from the endoplasmic reticulum (ER), intracellular trafficking and appearance in the plasma membrane, and removal via endocytosis play a key role in regulating Kv1 channel expression. In addition, interactions with specific intracellular enzymes can lead to the covalent modification of Kv1a and Kvb subunit polypeptides, impacting diverse characteristics of Kv1 channels in neuronal membranes. Many Kv1 channel a subunits carry specific intracellular trafficking signals that regulate their exit from the rough ER, where they are synthesized and inserted in a topologically correct manner into the membrane, and their stepwise transit through the biosynthetic endomembrane system. Polarized sorting of Kv1 channels to axon- or dendrite-directed cargo vesicles in the trans-Golgi network, targeted insertion at discrete sites within axonal or dendritic plasma membrane subdomains, and active retention at these sites to maintain discrete patterns of channel localization in the face of membrane fluidity are also mediated by discrete signals on Kv1 channel a subunits and dictate which channels are acting at which sites in neurons. Kvb subunits can also profoundly influence these characteristics. Mechanisms that regulate channel assembly, folding, and ER export of Kv1 channels, as well as mechanisms used to establish their correct subcellular localization, are beginning to be understood in detail. The ER export competence of membrane proteins seems to be determined by two major mechanisms: the chaperone-based quality control machinery that senses and acts on the folding state of proteins while in the ER and trafficking control machinery recognizing specific trafficking determinants. Whether these systems operate independently to control exit of proteins from

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the ER or are simply different faces of the same system is not clear (Mancias and Goldberg, 2005). Specific mechanisms also exist to sort newly synthesized membrane proteins into axon- or dendrite-destined transport vesicles (Tang, 2001). It seems likely that, given the highly restricted localization of many Kv1 channels, mechanisms exist for insertion of channel transporting vesicles at specific sites (juxtaparanodal, axon initial segment, etc.). Once present in the plasma membrane, such restricted localization, as well as inclusion of Kv1 channels in high-density clusters, must be maintained in the face of the dynamic lateral mobility of the lipid bilayer (Hedstrom and Rasband, 2006). This chapter focuses on recent insights into the expression and localization of mammalian brain Kv1 channels, and the mechanisms mediating their intracellular trafficking and the generation and maintenance of their distinct subcellular localizations. This chapter focuses on Kv channels whose principal or a subunits are members of the classical Shaker-related mammalian Kv family (i.e., Kv1) and not other subfamilies of Kva subunits (i.e., Kv2–Kv11).

5.2 MOLECULAR PROPERTIES OF KV1 CHANNELS 5.2.1

Nomenclature of Kv1 Channel Subunits and Genes

Kv1 channels are members of a diverse gene family. The nomenclature system for Kv channel a subunits, originally proposed by Chandy and colleagues (Chandy, 1991; Gutman et al., 2005) and now widely accepted, is based primarily on the relatedness of amino acid sequences between the different Kva subunits. The principal physiologically permeant ion of these channels is denoted by the chemical symbol (K for potassium), followed by the abbreviation of the ligand, which in this case is voltage (V). The remainder of the nomenclature relates to the gene families within these ion channel groups. The prototypical Kv channels have been divided into four families (Kv1–Kv4) based on the relative similarity of their amino acid sequences and on their relatedness to their single-gene orthologues in Drosophila: Kv1 (Shaker), Kv2 (Shab), Kv3 (Shaw), and Kv4 (Shal). A parallel nomenclature for Kv channel a subunit genes has also been developed, with genes named KCN*, the four gene families assigned the letters A–D (i.e., Kv1–Kv4 ¼ KCNA–KCND), and the specific gene numbers following the Kv nomenclature (Kv1.1 ¼ KCNA1, Kv1.4 ¼ KCNA4, Kv2.1 ¼ KCNB1, etc.). This review focuses exclusively on mammalian Kv1a subunits encoded by paralogous KCNA genes that are the orthologues of the Drosophila Shaker gene (Salkoff et al., 1992). In mammals, there are seven different Kv1a subunits (Kv1.1–Kv1.7) that are the products of the KCNA1–KCNA7 genes, respectively. Only Kv1.1–Kv1.6 are expressed in mammalian brain (Kalman et al., 1998; Gutman et al., 2005). None of the Kv1a subunit genes generates messenger RNAs that are subject to alternative splicing (Chandy and Gutman, 1995). Kvb auxiliary subunits that coassemble with Kv1a subunits to form native mammalian brain Kv1 channels are also members of a family of related genes. There exist three genes encoding these Kv1-associated Kvb subunits: KCNAB1

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(Kvb1), KCNAB2 (Kvb2), and KCNAB3 (Kvb3). Alternative splicing of transcripts from the KCNAB1 gene yields three different Kvb1 subunits: Kvb1.1, Kvb1.2, and Kvb1.3. There are two alternative forms of Kvb2 (Kvb2.1 and Kvb2.2) generated by alternative splicing of transcripts from the KCNAB2 gene (Akhtar et al., 1999). These paralogous genes are the orthologues of the Drosophila Hyperkinetic gene (Chouinard et al., 1995). 5.2.2

Kv1a and Kvb Subunit Expression in Mammalian Brain

In mammalian brain, the expression of Kv1a and Kvb subunits is primarily restricted to neurons, although glial cells may also express a subset of the neuronal repertoire (Vacher et al., 2006). In neurons, Kv1 channels are found predominantly on axons and nerve terminals, although dendritic expression is also found in certain neurons (Trimmer and Rhodes, 2004). Kv1 family members exhibit extensive coassembly to generate heteromeric channels with distinct characteristics (Pongs, 1999). In addition, assembly with auxiliary subunits can dramatically impact expression, localization, and function of the resultant channel complexes (Li et al., 2006). For example, inclusion of the Kvb1.1 subunit in Kv1 channel complexes containing Kv1.1 or Kv1.2 dramatically alters channel gating properties, converting the channels from sustained, or delayed rectifier type, to rapidly inactivating, or A-type (Rettig et al., 1994). The specific subunit composition of native complexes can also dramatically impact the expression level, localization, and function of Kv1 channels in mammalian neurons (Vacher et al., 2006). The predominant Kv1 cellular staining pattern throughout the brain is neuronal and subcellularly axonal. The three most abundant Kv1 subunits expressed in mammalian brain, Kv1.1, Kv1.2, and Kv1.4, are found predominantly localized to axons and nerve terminals (Trimmer and Rhodes, 2004). In many cases, these subunits are components of heteromeric channel complexes, as Kv1.1, Kv1.2, and Kv1.4 exhibit precise patterns of colocalization (Rhodes et al., 1997) and extensive association as shown by copurification (Rhodes et al., 1997; Shamotienko et al., 1997; Coleman et al., 1999). However, the subunit composition of channels containing these subunits varies across brain regions (Scott et al., 1994a). The Kv1.1a subunit in mammalian brain appears to be segregated into two major subpopulations: one associated with Kv1.2 and the other associated with Kv1.4. Kv1.1 and Kv1.2 are found in the absence of Kv1.4 in cerebellar basket cell terminals (Fig. 5.1a) (McNamara et al., 1993; Wang et al., 1993, 1994; Rhodes et al., 1995, 1997; Laube et al., 1996; McNamara et al., 1996), in the juxtaparanodal membrane adjacent to axonal nodes of Ranvier (Fig. 5.1b) (Wang et al., 1993, 1994; Rhodes et al., 1995, 1997; Rasband et al., 1998, 1999; Vabnick et al., 1999; Rasband et al., 2002, Rasband, 2004), and in the terminal segments of axons (Dodson et al., 2003). Kv1.1 and Kv1.2 are also present at axon initial segments (Dodson et al., 2002; Inda et al., 2006; Van Wart et al., 2007), sometimes in association with Kv1.4 (Fig. 5.1c), where they control axonal action potential waveform and synaptic efficacy (Kole et al., 2007). Kv1.1/Kv1.2 channels also play a role in m-opioid receptor-mediated modulation of GABAergic inputs into basolateral amygdala neurons (Finnegan et al., 2006) and in serotonin-modulated glutamate release from thalamocortical nerve

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FIGURE 5.1 Immunofluorescence staining of Kv1a and Kvb subunits in mammalian brain. (a) Staining for Kv1.1 (blue), Kv1.2 (green), and Kvb2 (red) in adult rat cerebellar cortex. Note overlap of all three fluors yielding white signal in basket cell terminals (arrowheads). (b) Staining for Kv1.2 (blue) in adult rat optic nerve (green ¼ Nav channels, red ¼ Caspr). Image courtesy of Dr. Matthew Rasband. (c) Staining for Kv1.4 (green) in the axon initial segment of a cultured hippocampal neuron (blue ¼ MAP2). Image courtesy of Drs. Yasuhiro Ogawa and Matthew Rasband. (d) Kv1.2 (red) in adult rat hippocampus (green ¼ Kv2.1). Arrowheads point to prominent Kv1.2 staining in perforant path presynaptic terminals in the middle molecular layer of the dentate gyrus. (See the color version of this figure in Color Plate section.)

terminals (Lambe and Aghajanian, 2001). Low-threshold, slowly inactivating axonal Kv1.2-containing channels, presumably containing either Kv1.4a or Kvb1 subunits to confer inactivation, are involved in the flexible properties of intracortical axons of layer 5 pyramidal neurons and may contribute significantly to intracortical processing (Shu et al., 2007). Kv1-containing channels are also important in setting the firing rate of layer II/III pyramidal neurons (Guan et al., 2007). Kv1.1 and Kv1.4 are found robustly expressed in the relative absence of Kv1.2 within the striatal efferents in globus pallidus and pars reticulata of substantia nigra (Sheng et al., 1992; Rhodes et al., 1997). Within the excitatory circuitry of the hippocampus, a number of patterns for expression of these three Kv1a subunits emerge, providing a striking example of the complex heterogeneity of subunit association (Rhodes et al., 1997). Kv1.1, Kv1.2, and Kv1.4 are highly expressed in the middle third of the molecular layer of the dentate gyrus (Fig. 5.1d), where they are associated with axons and terminals of the medial perforant path (Sheng et al., 1992, 1994; Wang et al., 1993, 1994; Rhodes et al., 1995, 1997; Veh et al., 1995; Monaghan et al., 2001). Kv1.1, Kv1.2, and Kv1.4 are also found in Schaffer collateral axons, while Kv1.1 and Kv1.4 colocalize, in the absence of Kv1.2, in mossy fiber axons (Sheng et al., 1992, 1994; Wang et al., 1993, 1994; Cooper et al., 1998), where they regulate Ca2þ influx and transmitter release (Geiger and Jonas, 2000).

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However, in spite of their colocalization, it is not clear that heteromeric channel complexes containing coassociated Kv1.1, Kv1.2, and Kv1.4 are present on perforant path and Schaffer collateral axons. Lesions in entorhinal cortex have distinct effects on the distribution of Kv1.2 and Kv1.4 in the middle third of the dentate molecular layer, suggesting that while these subunits may colocalize at the light microscope level, they may be expressed, along with Kv1.1, on different components of the perforant path (Monaghan et al., 2001). Similar results are obtained with lesions placed in other subfields. For example, predominant Kv1 channels appear to be composed of Kv1.1 with Kv1.4 in CA3, while in the Schaffer collateral pathway Kv1.1 is likely to be associated with Kv1.2 and Kv1.4 (Monaghan et al., 2001). Electron microscopic immunohistochemical studies have demonstrated that Kv1.1, Kv1.2, and Kv1.4 are concentrated in the axonal membrane immediately preceding or within axon terminals (Wang et al., 1993, 1994; Cooper et al., 1998). The immunoreactivity for Kv1.1 and Kv1.2 has been localized to the preterminal axonal membrane in stratum radiatum (Wang et al., 1993, 1994), while immunoreactivity for Kv1.4 has been localized to the preterminal extensions of mossy fiber axons (Cooper et al., 1998). Activation of Kv1 channels at these sites can play a critical role in regulating nerve terminal excitability and thereby regulate neurotransmitter release, as shown by pharmacological and genetic knockdown of Kv1 function (Dodson and Forsythe, 2004). The other Kv1a subunits appear to be expressed at lower levels in mammalian brain. Kv1.6 is found predominantly in interneurons, although some dendritic staining is seen throughout the brain on principal cell dendrites, which also express Kv1.1 and Kv1.2 (Rhodes et al., 1997) and presumably underlie the “D” current (Storm, 2000; Yuan and Chen, 2006). Kv1.3 is highly expressed in the cerebellar cortex. The bulk of this expression is in the parallel fiber axons of cerebellar granule cells, as strong in situ hybridization signal is present in the granule cell layer (Kues and Wunder, 1992), while strong immunostaining (Veh et al., 1995) and 125Imargatoxin binding (specific for Kv1.2 and Kv1.3) (Koch et al., 1997) are found in the molecular layer. The molecular layer also contains high levels of staining for Kv1.1 (Fig. 5.1a) (Veh et al., 1995; Rhodes et al., 1997), suggesting that Kv1.1 and Kv1.3 have the opportunity to form heteromeric channels on parallel fibers. The expression of Kv1.5 in the brain is overall quite low (Felix et al., 1999). What Kv1.5 expression there is may be restricted to nonneuronal cells. For example, Kv1.5 and Kv1.3 are components of delayed rectifier currents in glia (Khanna et al., 2001b; Chittajallu et al., 2002; Pannasch et al., 2006) and endothelial cells (Millar et al., 2007). In mammals, Kv1.7 is expressed in skeletal muscle, heart, and pancreatic islets, but not brain (Kalman et al., 1998). In situ hybridization, immunoprecipitation, and immunohistochemical analyses have also localized sites of expression of Kvb1 and Kvb2 in mammalian brain (Rettig et al., 1994; Rhodes et al., 1995, 1996, 1997). Kvb2 appears to be a component of many, if not all, Kv1-containing channel complexes in mammalian brain, and immunoreactivity for Kvb2 is present in each and every location where immunoreactivity for Kv1-family a subunits is observed (Rhodes et al., 1996, 1997). The Kvb1 subunit, which exerts dramatic effects on the inactivation kinetics of Kv1

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channels, appears to be included more selectively. Interestingly, the pattern of immunoreactivity for Kvb1 closely matches the expression pattern for Kv1.1 and Kv1.4, in that Kvb1 is found to colocalize with Kv1.4 in the medial perforant path, in the mossy fiber pathway, and in striatal efferents to the globus pallidus (Rhodes et al., 1996, 1997). Kvb2 is found in the absence of Kvb1 at many sites that exhibit colocalized Kv1.1 and Kv1.2, for example, in cerebellar basket cell terminals (Fig. 5.1a) and juxtaparanodes of nodes of Ranvier (Rhodes et al., 1997). As such there exist distinct pairings of Kv1a and Kvb subunits in heteromeric channel complexes in mammalian brain. 5.2.3

Structure of Kv1 Subunits

Detailed information on the primary, secondary, tertiary, and quaternary structures of mammalian Kv1a subunits is now available. Kv1a subunits exhibit extensive amino acid sequence identity within the transmembrane and pore-forming domains. The Shaker Kv channel from Drosophila melanogaster was the first Kv channel to be characterized (Tempel et al., 1987) and remains one of the best-studied Kv channels. Mammalian Kv1a subunits exhibit
70% amino acid identity with Shaker, and only 50% identity with a subunits from other mammalian Kv channel subfamilies Kv2– Kv4 (Salkoff et al., 1992). Shaker channels (MacKinnon, 1991) and mammalian Kv1 channels (Long et al., 2005) are tetramers of a subunits, and each a subunit consists of six transmembrane segments S1–S6 (Fig. 5.2a). The fourth transmembrane segment, or S4, of each subunit acts as the main component of the “voltage sensor” module, which detects the electrical potential across the membrane and controls the voltage-dependent gating of the channel (Papazian et al., 1991; Yellen, 1998; Swartz, 2004). The ionic conductance pathway or pore, which is responsible for rapid and selective potassium ion flux, is formed by the close association of the last two transmembrane segments (S5 and S6) from each of the four a subunits around a central water-filled cavity (Doyle et al., 1998; Long et al., 2005). Kvb subunits exhibit weak overall sequence similarity (McCormack and McCormack, 1994) but striking structural similarity (Gulbis et al., 1999) to aldo-keto reductase enzymes, and enzymatic activity against artificial substrates has recently been demonstrated (Weng et al., 2006). The structure (Fig. 5.2b) of a mammalian Kv1 channel complex formed by four Kv1.2a and four Kvb2 subunits reveals striking structural details of a Kv1 channel complex (Long et al., 2005). The portion of the Kv1.2a subunit between transmembrane segments S1–S4 comprises the voltage-sensing module, which is a distinct structure from the pore-forming module comprising the portion of the Kv1.2a subunit between transmembrane segments S5–S6 (Long et al., 2005). The modular structure of these domains and the relatively independent nature of the voltage sensor module are consistent with recent functional studies (Soler-Llavina et al., 2006). However, the specific conformation of the voltage sensor and pore modules in their resting and active states, and how the voltage-dependent movement of one is allosterically coupled to opening of the pore, remains unresolved and a focus of much recent research (Swartz, 2004).

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FIGURE 5.2 Structure of Kv1a and Kvb subunits. (a) Cartoon of the predicted topology of Kv1a subunits. The predicted transmembrane topology of a Kv1a subunit and sites defined as important determinants of intracellular trafficking and polarized expression are shown schematically. T1, tetramerization domain important for Kvb-mediated intracellular and polarized trafficking of Kv1a subunits; NLG, N-linked glycosylation site of Kv1.1–Kv1.5; ERR, ER retention motif found in Kv1.1, Kv1.2, and Kv1.6; FTS, forward trafficking signal in Kv1.4; PDZ BD, PDZ-binding motif in all Kv1a subunits; Y132, tyrosine residue critical for tyrosine kinasedependent suppression of Kv1.2; S440/S441, serine residues critical for phosphorylationdependent trafficking of Kv1.2. (b) Crystal structure of the Kv1.2/Kvb2 complex (Protein Data Bank accession number 2A79). Only two Kv1.2a and Kvb2 subunits are shown for clarity. TM, transmembrane domain; T1, tetramerization domain; Kvb, Kvb2 subunit. (c) Higher resolution view of the pore region of Kv1.2 with residues critical to the function of the ERR motif highlighted. Views of the Kv1.2 pore helices (S5 and S6) in blue with the side chains of the four critical residues depicted. The top panel shows wild-type Kv1.2 and the bottom panel the predicted structure of Kv1.2 with the four residues critical to the ERR function of Kv1.1 substituted at the equivalent positions. Note the change in the nature of the side chains forming the external face of the channel. Figures generated with Swiss PDB viewer. (See the color version of this figure in Color Plate section.)

5.3 DETERMINANTS OF INTRACELLULAR TRAFFICKING OF KV CHANNELS 5.3.1

Trafficking Signals Regulating Retention of Kv1 Channels in the ER

Neurons use diverse mechanisms to control the cell surface expression levels of Kv1 channels, which can profoundly affect neuronal excitability and signaling (Hille, 2001). Different levels of expression of Kv1a subunits lead, by mass action, to their proportional incorporation into channel complexes and the formation of tetrameric channels with different stoichiometries of Kv1a subunits. The precise stoichiometry of Kv1a subunits within a tetramer influences channel trafficking, localization, and function. Kv1 homomeric channels formed by different a subunits exhibit distinct protein stabilities and internal organelle retention characteristics. These patterns are altered upon coassembly to heteromeric complexes, such that the precise composition of Kv1a and Kvb subunits could affect both the overall level of the protein and its

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trafficking characteristics. As such, subunit composition can affect Kv1 channel cell surface expression levels as well as function. The overall steady-state cell surface expression levels and subunit composition appear to be regulated by a hierarchical system of regulatory steps, many of which operate at the level of the ER. The ER is a highly versatile protein factory that is equipped with chaperones and folding enzymes essential not only for protein folding and exit but also for retention and degradation of misfolded or aberrantly assembled protein complexes. Different Kv1a subunits exhibit high amino acid sequence identity (Stuhmer et al., 1989) but show striking differences in trafficking and functioning (Papazian, 1999; Manganas and Trimmer, 2000; Trimmer and Rhodes, 2004). The primary determinant for regulating trafficking of Kv1a subunits appears to be a potent ER retention (ERR) signal consisting of residues in the turret region (Fig. 5.2a) at the external face of the channel pore domain (Manganas et al., 2001b; Zhu et al., 2001). Cell surface expression of Kv1 subunits can also be influenced by a cytoplasmic C-terminal VXXSL motif (Fig. 5.2a) that acts as a forward trafficking signal (FTS) (Li et al., 2000) and by auxiliary Kvb subunits (Shi et al., 1996). In heterologous cells, these motifs dictate the steady-state distribution of Kv1 channels, such that homotetrameric Kv1.1 (and Kv1.6) channels are essentially localized to the ER (Fig. 5.3a), Kv1.4 channels mainly to the cell surface (Fig. 5.3b), and Kv1.2 channels to both the ER (Fig. 5.3c) and

FIGURE 5.3 Subcellular localization of homotetrameric Kv1a subunits in COS-1 cells. Recombinant Kv1 expression in monkey kidney fibroblast COS-1 cells. Intact cells were stained with ectodomain-directed antibodies (red), and then detergent permeabilized and stained with cytoplasmic domain-directed antibodies (green). (a) Kv1.1, (b) Kv1.4, (c and d) Kv1.2. (See the color version of this figure in Color Plate section.)

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the cell surface (Fig. 5.3d) (Manganas and Trimmer, 2000; Tiffany et al., 2000; Zhu et al., 2003). Kv1.1 homomeric channels in the ER appeared to be properly folded and assembled as tetramers with no evidence of aggregation or gross misfolding (Manganas et al., 2001a; Zhu et al., 2003). The generation of chimeras between Kv1.1 and Kv1.4 revealed that any Kv1.4a subunit containing the Kv1.1 pore region (P-loop), including the turret domain, was ER retained (Manganas et al., 2001b). Conversely, any Kv1.1a subunit containing the Kv1.4 P-loop was efficiently exported from the ER. An alignment of the Kv1.1 and Kv1.4 P-loop sequences revealed three key positions, all in the turret domain, which differed between Kv1.1 and Kv1.4 (Fig. 5.2a). Mutation of those in Kv1.1 to those in Kv1.4 yielded functional Kv1 channels that have Kv1.4-like trafficking patterns, and vice versa (Manganas et al., 2001b). Studies of these determinants for Kv1 channel surface expression have been primarily conducted in heterologous expression systems, although critical experiments have been reproduced in mammalian neurons, where the Kv1.1 ERR signal was also shown to function in cultured hippocampal neurons (Manganas et al., 2001b). Analyses of a large number of Kv1.1/Kv1.4 chimeras and truncation, deletion, and point mutants suggest that this luminal ERR motif is dominant over the cytoplasmic Cterminal FTS and also dominant over any effects of Kvb subunit coexpression on Kv1a subunit trafficking (Manganas et al., 2001b). Although small but measurable increases of Kv1.1 surface expression have been observed in cells coexpressing Kvb2 subunits (Shi et al., 1996), the effects are small compared to the effects of Kvb auxiliary subunits on homotetrameric Kv1.2 channels (Shi et al., 1996; Campomanes et al., 2002). Three amino acid residues (A352, E353, and Y379) within the highly conserved pore region of Kv1.1 are critical to functioning of the ERR (Manganas et al., 2001b). Subsequent studies yielded similar results and also revealed an additional role for S369 in the Kv1.1 ERR (Zhu et al., 2003, 2005). The function of the Kv1.1 ERR appears to be dictated only by these four pore regions of amino acid residues (Fig. 5.2c), in that point mutations in other Kv1 subunits at equivalent positions control the cell surface trafficking efficiency of these a subunits (Manganas et al., 2001b; Zhu et al., 2005). However, regulation of trafficking via these positions in the pore region is not transferable to Kv2, Kv3, and Kv4 subfamily members and seems to be unique to the Kv1 subfamily (Zhu et al., 2005). The location of the pore-localized ERR signal in Kv1.1 and the equivalent positions that regulate trafficking of other Kv1a subunits are unusual among membrane proteins in their ER luminal localization. One intriguing and surprising aspect of these findings is that three of the four P-loop residues (A352, E353, and Y379 in Kv1.1) that dictate Kv1 family trafficking also determine high-affinity binding to the mamba snake neurotoxin a-dendrotoxin or a-DTX (Hurst et al., 1991; Tytgat et al., 1995; Imredy and MacKinnon, 2000). Moreover, each of the Kv1 family members that binds a-DTX (Kv1.1, Kv1.2, and Kv1.6) exhibit a strong degree of ERR relative to the Kv1 family members (Kv1.3, Kv1.4, and Kv1.5) that lack the critical binding residues and do not bind a-DTX (Dolly and Parcej, 1996; Manganas and Trimmer, 2000; Tiffany et al., 2000; Manganas et al., 2001b). This allows for speculation that the ERR function of Kv1.1 (and of Kv1.2 and Kv1.6) may be mediated by a resident ERR receptor protein that binds to the turret domain of Kv1.1 in a fashion similar to the binding of a-DTX. Such a model is

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supported by the fact that exogenous expression of soluble DTX in the lumen of the ER leads to release of Kv1.1 from ER retention and efficient expression of Kv1.1 homotetrameric channels on the cell surface (Vacher et al., 2007). This mechanism is attractive in that should the resident ERR receptor resembles a-DTX, it may block the pore of ER-retained Kv1 channels, preventing any deleterious effects of ERlocalized Kv1 channels on the ionic integrity of the ER. This hypothetical ERR receptor for Kv1.1 and other a-DTX-sensitive Kv1a subunits remains to be identified. 5.3.2

Trafficking Signals Responsible for ER Export

Intrinsic sequences and/or interaction with auxiliary subunits can regulate ER export. Kv1.4 homotetrameric channels expressed in a wide variety of heterologous expression systems (COS-1, HEK293, CHO, MDCK cell lines), and in cultured hippocampal neurons, are efficiently expressed on the cell surface (Bekele-Arcuri et al., 1996; Shi and Trimmer, 1999; Li et al., 2000; Manganas and Trimmer, 2000). A large proportion of the steady-state Kv1.4 cellular pool (80–90%) is present on the cell surface (Fig. 5.3b). This is due to the lack of the critical amino acids in the turret region that mediate ER retention in Kv1.1 (Manganas et al., 2001b), combined with the presence of a unique ER export or FTS in the cytoplasmic carboxyl-terminal region (Li et al., 2000). This cytoplasmic FTS appears to be recessive to the turret domain ERR signal (which would be luminal in the ER), as Kv1.4 chimeras with active ERR signals from Kv1.1 but that are otherwise composed of Kv1.4 (including an intact FTS) are efficiently retained in the ER. However, simple deletion of the FTS decreases the level of Kv1.4 surface expression (Li et al., 2000; Zhu et al., 2003). Kvb subunits do not normally affect the trafficking of wild-type Kv1.4a subunits. This may be simply due to the fact that the inherent trafficking properties of these proteins may be near the maximum efficiency possible. However, Kv1.4 mutants lacking the FTS exhibit robust responses to Kvb coexpression (Li et al., 2000). Kvb1 and Kvb2 coexpression promotes the stability, N-linked glycosylation, and surface expression of homomeric Kv1.2 channels (Shi et al., 1996; Campomanes et al., 2002). The effects of Kvb subunits such as Kvb2 are mainly to increase the overall levels of those subunit combinations that already exhibit intermediate trafficking efficiencies, in that Kvb2 effects are obvious for Kv1.2, heteromeric channels containing Kv1.1, Kv1.2, and Kv1.4, and FTS-deficient Kv1.4 mutants, but are not detectable for Kv1.1 or Kv1.4 homotetramers. One possible explanation is that the main consequence of Kvb2 association is the stabilization of Kv1a subunits in the ER, as observed for Kv1.2 homotetramers (Shi et al., 1996). This stabilization could indirectly impact surface abundance by increasing the number of chances that a Kv1 channel complex would have to achieve ER export competence, through chaperone-assisted folding and/or saturation of retention receptor sites, by simply increasing the half-life of the channel complex in the ER. The fact that Kvb subunits such as Kvb2 are able to similarly promote plasma membrane abundance of different Kv1 channels regardless of precise a subunit composition implies that these cytoplasmic subunits are not directly involved in masking retention signals, the mechanism used by KChIP cytoplasmic subunits to facilitate trafficking of Kv4

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channels (Shibata et al., 2003; Scannevin et al., 2004), but mediate their effects indirectly via other mechanisms, such as stabilization or folding. 5.3.3

Posttranslational Regulation of Kv1 Channel Expression Levels

Among the enormous variety of posttranslational modifications, the only known modifications of Kv1a subunits are asparagine N-linked glycosylation (NLG) (Shi and Trimmer, 1999) and phosphorylation (Yang et al., 2007), and for Kvb subunits only phosphorylation (Gong et al., 1999). Posttranslational modification of proteins can play an important role in the proper folding, assembly, and trafficking, as well as in dynamic regulation of function. In this section, we will discuss the role of glycosylation and phosphorylation of Kv channels in channel trafficking. NLGs of membrane and secreted proteins have been shown to promote proper protein folding in the ER as well as to alter protein transport and targeting (Helenius and Aebi, 2001). In general, NLGs of Kv1 channels are not required for efficient trafficking to the cell surface but do appear to increase their surface expression by decreasing channel turnover rates and increasing channel stability. For example, blocking NLG of Shaker-type Kv channels dramatically decreases channel stability and steady-state cell surface expression levels but does not affect the folding (Santacruz-Toloza et al., 1994) and assembly of functional channels, or their transport to the cell surface (Khanna et al., 2001a). However, overexpression of the lectin-like protein chaperone calnexin (Bergeron et al., 1994; Hammond and Helenius, 1994) acts to promote Kv1.2 trafficking and increased cell surface expression, presumably through an interaction with NLGs (Manganas and Trimmer, 2004). Kv1.1–Kv1.5, but not Kv1.6, a subunits contain a single consensus site for NLG, which is located in the extracellular linker segment between transmembrane segments S1 and S2 (Fig. 5.2a). In native mammalian brain Kv1.1, Kv1.2, and Kv1.4a subunits, this NLG site carries a sialic acid-bearing oligosaccharide chain, conferring a strong negative charge to the extracellular glycan chain (Sheng et al., 1993; Thornhill et al., 1996; Shi and Trimmer, 1999; Manganas and Trimmer, 2000). Absence of these negative charges affects channel function (Thornhill et al., 1996; Watanabe et al., 2003). Important to many of the published studies of Kv1 trafficking, the NLG of these Kv1a subunits, and the spatial segregation of oligosaccharide-processing enzymes within the secretory pathway, allows tracking of the biosynthetic trafficking of Kv1a subunits by straightforward biochemical analyses of the glycan chain (Trimmer, 1998). Kv1 channel a subunits are also extensively modified by covalent phosphorylation. This posttranslational modification, governed by a diverse array of protein kinases and phosphatases, can modulate the expression and the function of supramolecular complexes formed from Kv1a subunits, Kvb auxiliary subunits, and interacting proteins. One important question is whether phosphorylation can dynamically regulate Kv1 trafficking. Steady-state Kv1.1 current density increases in transfected cells upon stimulation of specific protein kinases (Winklhofer et al., 2003). However, whether this is mediated through effects on biosynthetic trafficking, endocytosis/ turnover, or through changes in biophysical properties of active surface channels has not been directly assessed. Muscarinic stimulation of M1 receptors, or treatment with

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phorbol esters, tyrosine phosphatase inhibitors, and Ca2þ ionophore, yields enhanced Kv1.2 tyrosine phosphorylation and acute suppression of Kv1.2 functional expression (Huang et al., 1993; Tsai et al., 1997). Mutating a specific Tyr (Y132) in the Kv1.2 N-terminal linker connecting the T1 domain to the first S1 transmembrane segment (Fig. 5.2a) reduces the extent of stimulus-induced suppression by
50% (Huang et al., 1993). The bulk of the suppression of Kv1.2 functional expression in response to these treatments appears to be due to stimulus-induced increases in Kv1.2 endocytosis (Nesti et al., 2004). Kv1.2 interaction with the actin binding protein cortactin, which can also influence levels of Kv1.2 surface channels, is also regulated through tyrosine phosphorylation of Kv1.2 (Hattan et al., 2002). Expression of the Kv1.2a subunit appears to be especially sensitive to regulation by phosphorylation. Recent application of mass spectrometry-based identification of in vivo phosphorylation sites on ion channels (Park et al., 2008) has allowed for identification of novel serine phosphorylation sites on the C-terminus of Kv1.2a subunits purified from mammalian brain (Yang et al., 2007). Interestingly, analyses of Kv1.2 purified from rat, mouse, and human brain led to identification of the same set of serine phosphorylation sites, suggesting a conserved and fundamental function for phosphorylation at these C-terminal sites. As detailed above, Kv1.2a subunits are unique among Kv1a subunits in that they lack strong ERR (Manganas et al., 2001b) and FTS (Li et al., 2000) trafficking signals, such that homotetrameric Kv1.2 channels exhibit highly variable trafficking characteristics heavily influenced by other coexpressed Kv1a (Manganas and Trimmer, 2000) and auxiliary Kvb (Shi et al., 1996; Tiffany et al., 2000; Campomanes et al., 2002) subunits. Kv1.2 trafficking also differs between different cells in a population (Fig. 5.3c and d) (Shi et al., 1996; Manganas and Trimmer, 2000; Tiffany et al., 2000), and between different cell types (Manganas and Trimmer, 2000). This suggests conditional regulation of Kv1.2 trafficking, as can occur upon changes in protein phosphorylation state as recently shown for the KCNK3 potassium channel (O’Kelly et al., 2002). Kv1.2 phosphorylation at a subset of these sites (S440 and S441) identified in vivo occurs during intracellular trafficking of Kv1.2 (Fig. 5.2a), such that cell surface Kv1.2 channels are phosphorylated at these sites, but ER Kv1.2 pools are not. Mutation of these sites suppresses intracellular trafficking and leads to accumulation of Kv1.2 in the ER (Yang et al., 2007). Importantly, inclusion of Kv1.2 with altered phosphorylation state into heteromeric Kv1.2/Kv1.4 complexes also suppresses their trafficking, suggesting that incorporation of Kv1.2 into heteromeric Kv1 channel complexes such as predominate in brain may allow for dynamic regulation of their trafficking through stimulus-induced changes in phosphorylation state (Yang et al., 2007). The amplitude of Kv1.2 currents in heterologous cells is enhanced upon PKA activation (Huang et al., 1994). This involves phosphorylation at an N-terminal residue (T46), which leads to increased channel open probability and larger Kv1.2 currents (Huang et al., 1994). A recent report suggests that regulation of Kv1.2 by cAMP-based signaling is complex, with PKA-dependent and independent events (Connors et al., 2008). Such regulation involves not only T46 but also two of the C-terminal Ser sites identified as being phosphorylated in vivo in the studies detailed above (Yang et al., 2007). Changes in Kv1.2 phosphorylation appear to regulate cell surface expression

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levels and activity of Kv1.2 channels through effects on trafficking, endocytosis, and gating and may provide a crucial link between cell signaling pathways and electrical excitability. 5.3.4

Auxiliary Subunits of Kv1 Channels and Intracellular Trafficking

The best characterized of ion channel auxiliary subunits are the cytoplasmic Kvb subunits associated with Kv1 family members (Pongs et al., 1999). The bulk of Kv1 channel complexes in mammalian brain have associated Kvb subunits (Rhodes et al., 1996). Kvb subunits lack putative transmembrane domains and potential glycosylation sites, suggesting that they are cytoplasmic proteins (Scott et al., 1994b). Moreover, cryo-electron microscopy studies place their localization on the cytoplasmic face of the channel complex (Orlova et al., 2003; Sokolova et al., 2003). The Kvb subunits are each approximately 300 amino acids in length and share a common conserved core (over 85% amino acid identity), with the highest degree of variability in the amino terminus. The Kvb subunits are tetramers of oxidoreductase-like proteins arranged with fourfold symmetry similar to that of the integral membrane a subunits (Gulbis et al., 1999). Each oxidoreductase-like Kvb subunit contains an intact oxidoreductase active site with critical catalytic residues, and a bound NADPH (the reduced form of nicotinamide adenine dinucleotide phosphate) cofactor, but the specific substrate for any enzymatic activity is unknown, and the biological function of the oxidoreductase-like structure of Kvb subunits remains a mystery (Gulbis et al., 1999). Activity against artificial substrates has recently been demonstrated (Weng et al., 2006), a crucial first step toward identifying in vivo substrates. Studies of Kv1 channel biosynthesis have shown that Kv1a and Kvb subunits coassemble in the ER and remain together as a permanent complex (Shi et al., 1996; Nagaya and Papazian, 1997). Kvb subunits attach to a Kv1 channel through an interaction with the N-terminal cytoplasmic T1 domain (Fig. 5.2a and b) (Sewing et al., 1996; Yu et al., 1996; Gulbis et al., 2000; Long et al., 2005). Differences in the functional effects of Kvb subunits can often be ascribed to the variations in the amino terminal variable domain. Arguably, the most dramatic functional effect conferred by the association of Kvb subunits with Kv1 channels is an increase in the kinetics of channel inactivation gating (Heinemann et al., 1995). In a striking example, Kvb1.1 subunits can normally convert noninactivating delayed rectifier Kv channels to a rapidly inactivating channel (Rettig et al., 1994; Leicher et al., 1998). Kvb1 and Kvb2 have also been shown to modulate voltage dependence of activation of Kv1 channels in heterologous expression systems (England et al., 1995). A number of additional roles have been proposed for the function of Kvb/Kv1a subunit interactions. Each of the Kvb subunits can promote the cell surface expression of coexpressed Kv1.2a subunits (Shi et al., 1996; Campomanes et al., 2002). In these cases, Kvb subunits may aid in proper protein folding and/or a subunit assembly and thus enhance transport from ER to Golgi to cell surface. Effects of Kvb subunits on inactivation (Tipparaju et al., 2007), but not trafficking (Campomanes et al., 2002), can be regulated by intrinsic oxidoreductase activity. Recent studies showed that calnexin, the resident ER trafficking machinery, can also regulate Kv1 channel trafficking (Manganas and Trimmer, 2004). Calnexin is a

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type I integral membrane protein found in the ER that functions as a molecular chaperone to facilitate the folding and assembly of newly synthesized membrane proteins (Bergeron et al., 1994; Hammond and Helenius, 1994). Calnexin has been defined as a lectin chaperone that binds to membrane glycoproteins. The interaction occurs between the N-terminal ER luminal domain of the calnexin and the luminal NLGs (Ou et al., 1993). Homotetrameric Kv1.2 channels, possessing a weak porelocalized ERR (Manganas et al., 2001b) and FTS (Li et al., 2000) signals responded to calnexin coexpression with increased intracellular trafficking and cell surface expression (Manganas and Trimmer, 2004). This suggests that calnexin may be acting in its classical role to assist in efficient folding and/or increased stability of Kv1.2a subunit glycoprotein in the ER. Moreover, the interaction of the Kv1.2 N-terminus with Kvb2 subunits increases the efficiency of addition of the N-linked oligosaccharide chain of Kv1.2a subunits (Shi and Trimmer, 1999) through an unknown mechanism but that presumably occurs through an allosteric conformational change at or near the site of addition of the N-linked oligosaccharide chain that is induced by Kvb binding. This in itself may lead to a higher proportion of Kv1.2a subunits that are competent for interaction with calnexin and a subsequent increase in the number of channels that would be correctly folded and thus ER export competent. Kvb2 subunits and calnexin coexpression are epistatic, suggesting that they share a common pathway for promoting Kv1.2 channel surface expression (Manganas and Trimmer, 2004). In contrast, the trafficking of homotetrameric Kv1.1 channels, which contain strong ERR motifs (and no FTR export motifs), could not be influenced by calnexin coexpression (Manganas and Trimmer, 2004). These studies together demonstrate that a diverse set of intrinsic trafficking motifs, covalent modifications, and interacting proteins are crucial to intracellular trafficking of Kv1 channels.

5.4 DETERMINANTS OF SUBCELLULAR COMPARTMENTALIZATION OF KV1 CHANNELS IN THE PLASMA MEMBRANE Polarized localization of ion channels is essential for neurons to generate and maintain local and specialized signaling domains such as that found at synapses and the nodes of Ranvier. As discussed above, different Kv1 channels show distinct patterns of subcellular localization in neurons (Fig. 5.1). In this section, we discuss possible mechanisms underlying the polarized localization of Kv1 channels observed in neurons and its dynamic regulation. There are several hurdles to study intracellular trafficking of Kv channels compared to other ion channels, such as glutamate receptors: (1) Kv1 channels form complex heteromultimeric channels in neurons, (2) both the N- and C-termini are in the cytoplasm, and extracellular domains are limited to relatively short segments between transmembrane segments, which complicates analyses of surface pool of proteins by using epitope tags, biotinylation, or ectodomain-directed antibodies, and (3) expression levels of Kv1 channels are relatively low. Despite these complications, there are several compelling studies describing trafficking determinants of Kv1 channels that provide insights into molecular mechanisms determining polarized localization of Kv1 channels in central neurons.

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SHAKER FAMILY KV1 VOLTAGE-GATED POTASSIUM CHANNELS

Determinants of Axonal Localization of Kv1 Channels

The precise determinants for the polarized expression and local clustering of most Kv1 channels are not well characterized. One of the problems has been recapitulating the heteromeric channels found in native neurons in model cell systems (e.g., polarized epithelial cells), compounded by the inefficient intracellular trafficking inherent to many of the Kv1a subunits. Moreover, for reasons that are not well understood, efficiently expressed Kv1a subunits, such as Kv1.4, when expressed in such model systems do not “behave”; for example, axonal Kv1.4 is found in the basolateral and not apical membranes of polarized MDCK epithelial cells (Le Maout et al., 1996). One possible explanation for such discrepancies is that recent studies have provided compelling data that cytoplasmic Kvb subunits are crucial to polarized trafficking of Kv1a subunits. While Kv1.2a subunits overexpressed in cultured hippocampal neurons exhibit somatodendritic localization, cotransfection of the Kvb2 subunit yields a pronounced axonal localization of Kv1.2 (Campomanes et al., 2002). Subsequent analyses of Kv1.2 deletion mutants and chimeric channels revealed that the amino terminal T1 domain, which comprises the Kvb subunit binding site (Sewing et al., 1996; Yu et al., 1996; Gulbis et al., 2000), is essential for axonal expression of Kv1.2 channels (Gu et al., 2003). Specific mutations to disrupt Kvb2 binding also disrupted axonal expression of Kv1a subunits in transfected neurons, further suggesting that endogenous Kvb subunits were contributing to the axonal Kv1 localization (Gu et al., 2003). Remarkably, the wild-type Kv1.2 T1 domain, but not mutants with altered Kvb binding, was able to direct the axonal localization of single-pass transmembrane reporter proteins (Gu et al., 2003). Thus, cytoplasmic Kvb subunits may affect not only early biosynthetic processing events and ER export (Shi et al., 1996), but also axonal localization (Campomanes et al., 2002; Gu et al., 2003) of Kv1 channels. Recent studies have shown that axonal targeting of Kv1 channels is dependent on the direct interaction of Kvb2 with the microtubule plus-end-tracking protein EB1 (Gu et al., 2006). Analysis of Kvb2 deletion mutants revealed that the disruption of its association with EB1 dramatically decreases Kv1.2 axonal targeting (Gu et al., 2006). Suppression of endogenous EB1 expression using siRNA also impairs axonal targeting of endogenous Kv1 channels but not voltage-gated sodium channels (Gu et al., 2006), which appear to target to axons by distinct mechanisms and motifs (Garrido et al., 2003). Another recent study shows that Kv1 channel axonal targeting is dependent not only on EB1 and Kvb2 but also on the interaction of the Kv1a subunit T1 domain with the kinesin KIF5B (Rivera et al., 2007). The role of Kvb2, which also binds at the T1 domain, was not addressed in this study, although endogenous Kvb2 was presumably expressed in the neurons in the cultured cortical slices used in these studies (Rivera et al., 2007). As such, the T1 domain of Kv1a subunits mediates multiple protein–protein interactions, including tetrameric assembly of Kv1a subunits, association with Kvb subunits (and their EB1 binding partner), and interaction with KIF5B. The important role of Kvb2 is underscored by the finding that deletion of the human Kvb2 gene in chromosome 1p36 deletion syndrome is closely linked to epilepsy that is present in a subset of these patients (Heilstedt et al., 2001) and that Kvb2 null mice also exhibit seizures and cold swim-induced tremors similar to that

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observed in Kv1.1-null mice (McCormack et al., 2002). However, the discrepancy that Kv1.2 localization is minimally affected in the cerebellar basket cell synaptic terminals in Kvb2 null mice remains to be addressed (McCormack et al., 2002), although Kvb1 can substitute for Kvb2 in effects on Kv1 trafficking (Shi et al., 1996; Campomanes et al., 2002), and Kvb1/Kvb2 double knockout mice have more severe defects than either single knockout (Connor et al., 2005). 5.4.2

Determinants of Kv1 Channel Clustering

Membrane-associated guanylate kinases (MAGUKs) such as PSD-95, Chapsyn-110, SAP102, and SAP97 function as scaffolding molecules to promote coclustering of membrane receptors and ion channels (Sheng and Sala, 2001). Mutational and structural analyses showed that the PDZ domains of PSD-95 and other MAGUKs bind to the PDZ-binding motif (S/TxV) in the carboxyl-termini of all Kv1a subunits (Fig. 5.2a) (Kim et al., 1995; Doyle et al., 1996; Kim and Sheng, 1996; Tiffany et al., 2000). In most mammalian central neurons, MAGUKs are found at synapses, most prominently in the postsynaptic density (Sheng and Sala, 2001) and Kv1 channels on axons (Trimmer and Rhodes, 2004). However, MAGUKs are found as prominent components of Kv1 complexes purified from mammalian brain (Schulte et al., 2006). Examples where Kv1 channels and MAGUKs, in this case PSD-95, exhibit robust colocalization in neurons are in the colocalization of Kv1.1, Kv1.2, and Kvb2 with PSD-95 in cerebellar basket cells terminals (Kim et al., 1995; Laube et al., 1996) and at the juxtaparanodal regions of nodes of Ranvier (Rasband and Trimmer, 2001). Although mutation of the Drosophila discs large gene, a homologue of PSD-95, results in failure to localize Shaker potassium channels to the neuromuscular junction of flies, mutation of PSD-95 in mice does not lead to altered Kv1 channel localization (Rasband et al., 2002). At the level of the light microscope, the localization of Kv1.1 and Kv1.2 in basket cell terminals and juxtaparanodes is normal in transgenic mice expressing a truncated form of PSD-95 that is no longer expressed at these sites. While MAGUKs remain as attractive candidates for the interacting proteins that cluster Kv1 channels at discrete plasma membrane subdomains in axons and near nerve terminals, the specific binding partners mediating the clustering and their regulation remain to be elucidated.

5.5 PATHOLOGICAL ALTERATIONS IN KV1 CHANNEL TRAFFICKING AND DISTRIBUTION Kv1 channels are major regulators of the membrane excitability of neuronal nerve terminals (Dodson and Forsythe, 2004). Failures in controlling Kv1 channel expression, localization, and function may result in severe pathological hyperexcitability, leading to epilepsy and/or ataxia. As detailed above, neurons display wellorganized patterns of Kv1 channel expression and localization that is tightly regulated through diverse cellular mechanisms. Aberrations in these molecular events could affect neuronal excitability and lead to neuronal dysfunction as discussed below.

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SHAKER FAMILY KV1 VOLTAGE-GATED POTASSIUM CHANNELS

Episodic Ataxia

Episodic ataxia type-1 (EA-1) is a neurological syndrome that is characterized by continuous myokymia and cerebellar ataxia (Kullmann, 2002). EA-1 is caused by mutations in the Kv1.1 subunit that recessively affect individuals (Kullmann, 2002). Genetic linkage studies in EA-1 patients have identified at least 17 mutations in the Kv1.1 gene (Cannon, 2006; Jen et al., 2007). Most of the EA-1 mutations are missense mutations that lead to Kv1.1 channels with altered biophysical characteristics (Zerr et al., 1998; Maylie et al., 2002). However, the single EA-1 nonsense mutation that results in the truncation of the last 79 amino acid residues in the C-terminus (Eunson et al., 2000) causes loss of expression, misfolding, and resultant intracellular aggregation of Kv1 channels (Manganas et al., 2001a). Interestingly, the expression of this mutant also leads to intracellular aggregation and retention of coassembled wild-type Kv1a and Kvb subunits (Manganas et al., 2001a), suggesting effects beyond those on Kv1.1a subunits. These dominant negative properties of the EA-1 mutant Kv1.1 resemble in many aspects to those of the mutant CFTR protein encoded by the predominant mutation in cystic fibrosis patients. 5.5.2

Epilepsy

Epilepsy is a common neurological disorder and, in some cases, can be linked to channel dysfunction. To date, single-gene ion channel mutations are the predominant cause of rare Mendelian idiopathic epilepsy syndromes (Mulley et al., 2003; Cannon, 2006), of which two encode Kv channels (Kv7.2 and Kv7.3) outside of the family on which this review focuses. Experimental knockout of Kv1.1 channels in mice results in early postnatal epilepsy (Smart et al., 1998), and knockout of Kv1.2 also yields mice with epilepsy and a shortened life span (Brew et al., 2007). This suggests significant roles of Kv1 channels in the development of epilepsy. Kv1.1 knockout mice have an enlarged hippocampus (Persson et al., 2007) and exhibit hippocampal damage and reorganization (Wenzel et al., 2007). However, expression levels and localization of other Kv1 family members, auxiliary subunits, and interacting proteins are remarkably unaffected (Wenzel et al., 2007), in sharp contrast to findings in Kv4.2 knockout mice (Menegola and Trimmer, 2006). A spontaneous mutation in mouse Kv1.1 was identified in the megencephaly mouse (Petersson et al., 2003). The mutation leads to expression of a severely truncated form of Kv1.1 that is phenotypically similar to the EA-1 truncation mutant described above (Manganas et al., 2001a), in that it has trafficking deficiencies that can be transferred to heteromeric Kv1 channel complexes (Persson et al., 2005). Remarkably, a recent report showed that the phenotypes of mice with a voltage-gated calcium channel mutation that leads to neuronal hypoexcitability and absence epilepsy, and the hyperexcitable seizure phenotype of Kv1.1 knockout mice, can be eliminated by simply crossing the two mice (Glasscock et al., 2007). This highlights the role of precise regulation of neuronal excitability in maintaining normal brain function and the molecular complexity of such regulation.

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5.6 CONCLUDING REMARKS Given the crucial role of Kv1 channels in regulating the excitability of mammalian presynaptic nerve terminal, somata, and dendrites, a better understanding of the molecular and cellular mechanisms of their intracellular trafficking subcellular targeting is of great biological and biomedical significance. Such information will not only provide insights into mechanisms that generate the observed patterns of expression and discrete localization of Kv1 channels in precise functional domains of the axonal and somatodendritic membrane, but could also provide information crucial to development of highly selective drugs that target this important class of ion channels and modulate neuronal function. Moreover, it is likely to provide important information as to the etiology and pathophysiology of diseases of neuronal dysfunction such as ataxia and epilepsy.

ACKNOWLEDGMENT This work was supported by National Institutes of Health Grant NS34383 (J. S. T.).

REFERENCES Akhtar S, McIntosh P, Bryan-Sisneros A, Barratt L, Robertson B, Dolly JO, 1999. A functional spliced-variant of beta 2 subunit of Kv1 channels in C6 glioma cells and reactive astrocytes from rat lesioned cerebellum. Biochemistry 38: 16984–16992. Bekele-Arcuri Z, Matos MF, Manganas L, Strassle BW, Monaghan MM, Rhodes KJ, Trimmer JS, 1996. Generation and characterization of subtype-specific monoclonal antibodies to Kþ channel alpha- and beta-subunit polypeptides. Neuropharmacology 35: 851–865. Bergeron JJ, Brenner MB, Thomas DY, Williams DB, 1994. Calnexin: a membrane-bound chaperone of the endoplasmic reticulum. Trends Biochem Sci 19: 124–128. Brew HM, Gittelman JX, Silverstein RS, Hanks T, Demas VP, Robinson LC, Robbins CA, McKee-Johnson J, Chiu SY, Messing A, Tempel BL, 2007. Seizures and reduced lifespan in mice lacking the potassium channel subunit Kv1.2, but hypoexcitability and enlarged Kv1 currents in auditory neurons. J Neurophysiol. 98: 1501–1525. Campomanes CR, Carroll KI, Manganas LN, Hershberger ME, Gong B, Antonucci DE, Rhodes KJ, Trimmer JS, 2002. Kv beta subunit oxidoreductase activity and Kv1 potassium channel trafficking. J Biol Chem 277: 8298–8305. Cannon SC, 2006. Pathomechanisms in channelopathies of skeletal muscle and brain. Annu Rev Neurosci 29: 387–415. Chandy KG, 1991. Simplified gene nomenclature. Nature 352: 26. Chandy KG, Gutman GA, 1995. Voltage-gated potassium channel genes. In: North RA (Ed.), Ligand- and Voltage-gated Ion Channels, CRC Press, Boca Raton, pp. 1–71. Chittajallu R, Chen Y, Wang H, Yuan X, Ghiani CA, Heckman T, McBain CJ, Gallo V, 2002. Regulation of Kv1 subunit expression in oligodendrocyte progenitor cells and their role in G1/S phase progression of the cell cycle. Proc Natl Acad Sci USA 99: 2350–2355.

146

SHAKER FAMILY KV1 VOLTAGE-GATED POTASSIUM CHANNELS

Chouinard SW, Wilson GF, Schlimgen AK, Ganetzky B, 1995. A potassium channel beta subunit related to the aldo-keto reductase superfamily is encoded by the Drosophila hyperkinetic locus. Proc Natl Acad Sci USA 92: 6763–6767. Coleman SK, Newcombe J, Pryke J, Dolly JO, 1999. Subunit composition of Kv1 channels in human CNS. J Neurochem 73: 849–858. Connor JX, McCormack K, Pletsch A, Gaeta S, Ganetzky B, Chiu SY, Messing A, 2005. Genetic modifiers of the Kv beta2-null phenotype in mice. Genes Brain Behav 4: 77–88. Connors EC, Ballif BA, Morielli AD, 2008. Homeostatic regulation of Kv1.2 potassium channel trafficking by cyclic AMP. J Biol Chem 283:3445–3453. Cooper EC, Milroy A, Jan YN, Jan LY, Lowenstein DH, 1998. Presynaptic localization of Kv1.4-containing A-type potassium channels near excitatory synapses in the hippocampus. J Neurosci 18: 965–974. Dodson PD, Forsythe ID, 2004. Presynaptic Kþ channels: electrifying regulators of synaptic terminal excitability. Trends Neurosci 27: 210–217. Dodson PD, Barker MC, Forsythe ID, 2002. Two heteromeric Kv1 potassium channels differentially regulate action potential firing. J Neurosci 22: 6953–6961. Dodson PD, Billups B, Rusznak Z, Szucs G, Barker MC, Forsythe ID, 2003. Presynaptic rat Kv1.2 channels suppress synaptic terminal hyperexcitability following action potential invasion. J Physiol 550: 27–33. Dolly JO, Parcej DN, 1996. Molecular properties of voltage-gated Kþ channels. J Bioenerg Biomembr 28: 231–253. Doyle DA, Lee A, Lewis J, Kim E, Sheng M, MacKinnon R, 1996. Crystal structures of a complexed and peptide-free membrane protein- binding domain: molecular basis of peptide recognition by PDZ. Cell 85: 1067–1076. Doyle DA, Morais Cabral J, Pfuetzner RA, Kuo A, Gulbis JM, Cohen SL, Chait BT, MacKinnon R, 1998. The structure of the potassium channel: molecular basis of Kþ conduction and selectivity. Science 280: 69–77. England SK, Uebele VN, Kodali J, Bennett PB, Tamkun MM, 1995. A novel Kþ channel betasubunit (hKv beta 1.3) is produced via alternative mRNA splicing. J Biol Chem 270: 28531– 28534. Eunson LH, Rea R, Zuberi SM, Youroukos S, Panayiotopoulos CP, Liguori R, Avoni P, McWilliam RC, Stephenson JB, Hanna MG, Kullmann DM, Spauschus A, 2000. Clinical, genetic, and expression studies of mutations in the potassium channel gene KCNA1 reveal new phenotypic variability. Ann Neurol 48: 647–656. Felix JP, Bugianesi RM, Schmalhofer WA, Borris R, Goetz MA, Hensens OD, Bao JM, Kayser F, Parsons WH, Rupprecht K, Garcia ML, Kaczorowski GJ, Slaughter RS, 1999. Identification and biochemical characterization of a novel nortriterpene inhibitor of the human lymphocyte voltage-gated potassium channel, Kv1. 3. Biochemistry 38: 4922–4930. Finnegan TF, Chen SR, Pan HL, 2006. Mu opioid receptor activation inhibits GABAergic inputs to basolateral amygdala neurons through Kv1.1/1.2 channels. J Neurophysiol 95: 2032– 2041. Garrido JJ, Fernandes F, Moussif A, Fache MP, Giraud P, Dargent B, 2003. Dynamic compartmentalization of the voltage-gated sodium channels in axons. Biol Cell 95: 437–445. Geiger JRP, Jonas P, 2000. Dynamic control of presynaptic Ca2þ inflow by fast-inactivating Kþ channels in hippocampal mossy fiber boutons. Neuron 28: 927–939.

REFERENCES

147

Glasscock E, Qian J, Yoo JW, Noebels JL, 2007. Masking epilepsy by combining two epilepsy genes. Nat Neurosci 10: 1554–1558. Gong J, Xu J, Bezanilla M, van Huizen R, Derin R, Li M, 1999. Differential stimulation of PKC phosphorylation of potassium channels by ZIP1 and ZIP2. Science 285: 1565–1569. Gu C, Jan YN, Jan LY, 2003. A conserved domain in axonal targeting of Kv1 (Shaker) voltagegated potassium channels. Science 301: 646–649. Gu C, Zhou W, Puthenveedu MA, Xu M, Jan YN, Jan LY, 2006. The microtubule plus-end tracking protein EB1 is required for Kv1 voltage-gated Kþ channel axonal targeting. Neuron 52: 803–816. Guan D, Lee JC, Higgs MH, Spain WJ, Foehring RC, 2007. Functional roles of Kv1 channels in neocortical pyramidal neurons. J Neurophysiol 97: 1931–1940. Gulbis JM, Mann S, MacKinnon R, 1999. Structure of a voltage-dependent Kþ channel beta subunit. Cell 97: 943–952. Gulbis JM, Zhou M, Mann S, MacKinnon R, 2000. Structure of the cytoplasmic beta subunit-T1 assembly of voltage- dependent Kþ channels. Science 289: 123–127. Gutman GA, Chandy KG, Grissmer S, Lazdunski M, McKinnon D, Pardo LA, Robertson GA, Rudy B, Sanguinetti MC, Stuhmer W, Wang X, 2005. International Union of Pharmacology LIII. Nomenclature and molecular relationships of voltage-gated potassium channels. Pharmacol Rev 57: 473–508. Hammond C, Helenius A, 1994. Folding of VSV G protein: sequential interaction with BiP and calnexin. Science 266: 456–458. Hattan D, Nesti E, Cachero TG, Morielli AD, 2002. Tyrosine phosphorylation of Kv1.2 modulates its interaction with the actin-binding protein cortactin. J Biol Chem 277: 38596– 38606. Hedstrom KL, Rasband MN, 2006. Intrinsic and extrinsic determinants of ion channel localization in neurons. J Neurochem 98: 1345–1352. Heilstedt HA, Burgess DL, Anderson AE, Chedrawi A, Tharp B, Lee O, Kashork CD, Starkey DE, Wu YQ, Noebels JL, Shaffer LG, Shapira SK, 2001. Loss of the potassium channel betasubunit gene, KCNAB2, is associated with epilepsy in patients with 1p36 deletion syndrome. Epilepsia 42: 1103–1111. Heinemann SH, Rettig J, Wunder F, Pongs O, 1995. Molecular and functional characterization of a rat brain Kv beta 3 potassium channel subunit. FEBS Lett 377: 383–389. Helenius A, Aebi M, 2001. Intracellular functions of N-linked glycans. Science 291: 2364– 2369. Hille B, 2001. Ionic channels of excitable membranes, 3rd ed. Sinauer, Sunderland, MA. Huang XY, Morielli AD, Peralta EG, 1993. Tyrosine kinase-dependent suppression of a potassium channel by the G protein-coupled m1 muscarinic acetylcholine receptor. Cell 75: 1145–1156. Huang XY, Morielli AD, Peralta EG, 1994. Molecular basis of cardiac potassium channel stimulation by protein kinase A. Proc Natl Acad Sci USA 91: 624–628. Hurst RS, Busch AE, Kavanaugh MP, Osborne PB, North RA, Adelman JP, 1991. Identification of amino acid residues involved in dendrotoxin block of rat voltage-dependent potassium channels. Mol Pharmacol 40: 572–576. Imredy JP, MacKinnon R, 2000. Energetic and structural interactions between deltadendrotoxin and a voltage-gated potassium channel. J Mol Biol 296: 1283–1294.

148

SHAKER FAMILY KV1 VOLTAGE-GATED POTASSIUM CHANNELS

Inda MC, DeFelipe J, Munoz A, 2006. Voltage-gated ion channels in the axon initial segment of human cortical pyramidal cells and their relationship with chandelier cells. Proc Natl Acad Sci USA 103: 2920–2925. Jen JC, Graves TD, Hess EJ, Hanna MG, Griggs RC, Baloh RW, 2007. Primary episodic ataxias: diagnosis, pathogenesis and treatment. Brain 130: 2484–2493. Kalman K, Nguyen A, Tseng-Crank J, Dukes ID, Chandy G, Hustad CM, Copeland NG, Jenkins NA, Mohrenweiser H, Brandriff B, Cahalan M, Gutman GA, Chandy KG, 1998. Genomic organization, chromosomal localization, tissue distribution, and biophysical characterization of a novel mammalian Shaker-related voltage-gated potassium channel Kv1. 7. J Biol Chem 273: 5851–5857. Khanna R, Myers MP, Laine M, Papazian DM, 2001a. Glycosylation increases potassium channel stability and surface expression in mammalian cells. J Biol Chem 276: 34028– 34034. Khanna R, Roy L, Zhu X, Schlichter LC, 2001b. Kþ channels and the microglial respiratory burst. Am J Physiol Cell Physiol 280: C796–C806. Kim E, Sheng M, 1996. Differential Kþ channel clustering activity of PSD-95 and SAP97, two related membrane-associated putative guanylate kinases. Neuropharmacology 35: 993–1000. Kim E, Niethammer M, Rothschild A, Jan YN, Sheng M, 1995. Clustering of Shaker-type Kþ channels by interaction with a family of membrane-associated guanylate kinases. Nature 378: 85–88. Koch RO, Wanner SG, Koschak A, Hanner M, Schwarzer C, Kaczorowski GJ, Slaughter RS, Garcia ML, Knaus HG, 1997. Complex subunit assembly of neuronal voltage-gated Kþ channels. Basis for high-affinity toxin interactions and pharmacology. J Biol Chem 272: 27577–27581. Kole MH, Letzkus JJ, Stuart GJ, 2007. Axon initial segment Kv1 channels control axonal action potential waveform and synaptic efficacy. Neuron 55: 633–647. Kues WA, Wunder F, 1992. Heterogeneous expression patterns of mammalian potassium channel genes in developing and adult rat brain. Eur J Neurosci 4: 1296–1308. Kullmann DM, 2002. The neuronal channelopathies. Brain 125: 1177–1195. Lambe EK, Aghajanian GK, 2001. The role of Kv1.2-containing potassium channels in serotonin-induced glutamate release from thalamocortical terminals in rat frontal cortex. J Neurosci 21: 9955–9963. Laube G, Roper J, Pitt JC, Sewing S, Kistner U, Garner CC, Pongs O, Veh RW, 1996. Ultrastructural localization of Shaker-related potassium channel subunits and synapseassociated protein 90 to septate-like junctions in rat cerebellar Pinceaux. Brain Res Mol Brain Res 42: 51–61. Le Maout S, Sewing S, Coudrier E, Elalouf JM, Pongs O, Merot J, 1996. Polarized targeting of a shaker-like (A-type) Kþ-channel in the polarized epithelial cell line MDCK. Mol Membr Biol 13: 143–147. Leicher T, Bahring R, Isbrandt D, Pongs O, 1998. Coexpression of the KCNA3B gene product with Kv1. 5 leads to a novel A-type potassium channel. J Biol Chem 273: 35095–35101. Levitan IB, 2006. Signaling protein complexes associated with neuronal ion channels. Nat Neurosci 9: 305–310. Li D, Takimoto K, Levitan ES, 2000. Surface expression of Kv1 channels is governed by a C-terminal motif. J Biol Chem 275: 11597–11602.

REFERENCES

149

Li Y, Um SY, McDonald TV, 2006. Voltage-gated potassium channels: regulation by accessory subunits. Neuroscientist 12: 199–210. Llinas RR, 1988. The intrinsic electrophysiological properties of mammalian neurons: insights into central nervous system function. Science 242: 1654–1664. Long SB, Campbell EB, Mackinnon R, 2005. Crystal structure of a mammalian voltagedependent Shaker family Kþ channel. Science 309: 897–903. MacKinnon R, 1991. Determination of the subunit stoichiometry of a voltage-activated potassium channel. Nature 350: 232–235. Mancias JD, Goldberg J, 2005. Exiting the endoplasmic reticulum. Traffic 6: 278–285. Mandel G, McKinnon D, 1993. Molecular basis of neural-specific gene expression. Annu Rev Neurosci 16: 323–345. Manganas LN, Trimmer JS, 2000. Subunit composition determines Kv1 potassium channel surface expression. J Biol Chem 275: 29685–29693. Manganas LN, Trimmer JS, 2004. Calnexin regulates mammalian Kv1 channel trafficking. Biochem Biophys Res Commun 322: 577–584. Manganas LN, Akhtar S, Antonucci DE, Campomanes CR, Dolly JO, Trimmer JS, 2001a. Episodic ataxia type-1 mutations in the Kv1.1 potassium channel display distinct folding and intracellular trafficking properties. J Biol Chem 276: 49427–49434. Manganas LN, Wang Q, Scannevin RH, Antonucci DE, Rhodes KJ, Trimmer JS, 2001b. Identification of a trafficking determinant localized to the Kv1 potassium channel pore. Proc Natl Acad Sci USA 98: 14055–14059. Maylie B, Bissonnette E, Virk M, Adelman JP, Maylie JG, 2002. Episodic ataxia type 1 mutations in the human Kv1.1 potassium channel alter hKvbeta 1-induced N-type inactivation. J Neurosci 22: 4786–4793. McCormack T, McCormack K, 1994. Shaker Kþ channel beta subunits belong to an NAD(P)Hdependent oxidoreductase superfamily. Cell 79: 1133–1135. McCormack K, Connor JX, Zhou L, Ho LL, Ganetzky B, Chiu SY, Messing A, 2002. Genetic analysis of the mammalian Kþ channel beta subunit Kvbeta 2 (Kcnab2). J Biol Chem 277: 13219–13228. McNamara NM, Muniz ZM, Wilkin GP, Dolly JO, 1993. Prominent location of a Kþ channel containing the alpha subunit Kv1.2 in the basket cell nerve terminals of rat cerebellum. Neuroscience 57: 1039–1045. McNamara NM, Averill S, Wilkin GP, Dolly JO, Priestley JV, 1996. Ultrastructural localization of a voltage-gated Kþ channel alpha subunit (KV 1.2) in the rat cerebellum. Eur J Neurosci 8: 688–699. Menegola M, Trimmer JS, 2006. Unanticipated region- and cell-specific downregulation of individual KChIP auxiliary subunit isotypes in Kv4 .2 knock-out mouse brain. J Neurosci 26: 12137–12142. Millar ID, Wang S, Brown PD, Barrand MA, Hladky SB, 2007. Kv1 and Kir2 potassium channels are expressed in rat brain endothelial cells. Pflugers Arch. 456: 379–391. Mohapatra DP, Park KS, Trimmer JS, 2007. Dynamic regulation of the voltage-gated Kv2.1 potassium channel by multisite phosphorylation. Biochem Soc Trans 35: 1064–1068. Monaghan MM, Trimmer JS, Rhodes KJ, 2001. Experimental localization of Kv1 family voltage-gated Kþ channel alpha and beta subunits in rat hippocampal formation. J Neurosci 21: 5973–5983.

150

SHAKER FAMILY KV1 VOLTAGE-GATED POTASSIUM CHANNELS

Mulley JC, Scheffer IE, Petrou S, Berkovic SF, 2003. Channelopathies as a genetic cause of epilepsy. Curr Opin Neurol 16: 171–176. Nagaya N, Papazian DM, 1997. Potassium channel alpha and beta subunits assemble in the endoplasmic reticulum. J Biol Chem 272: 3022–3027. Nesti E, Everill B, Morielli AD, 2004. Endocytosis as a mechanism for tyrosine kinasedependent suppression of a voltage-gated potassium channel. Mol Biol Cell 15: 4073–4088. O’Kelly I, Butler MH, Zilberberg N, Goldstein SA, 2002. Forward transport 14-3-3 binding overcomes retention in endoplasmic reticulum by dibasic signals. Cell 111: 577–588. Orlova EV, Papakosta M, Booy FP, van Heel M, Dolly JO, 2003. Voltage-gated Kþ channel from mammalian brain: 3D structure at 18A of the complete (alpha)4(beta)4 complex. J Mol Biol 326: 1005–1012. Ou WJ, Cameron PH, Thomas DY, Bergeron JJ, 1993. Association of folding intermediates of glycoproteins with calnexin during protein maturation. Nature 364: 771–776. Pannasch U, Farber K, Nolte C, Blonski M, Yan Chiu S, Messing A, Kettenmann H, 2006. The potassium channels Kv1.5 and Kv1. 3 modulate distinct functions of microglia. Mol Cell Neurosci 33: 401–411. Papazian DM, 1999. Potassium channels: some assembly required. Neuron 23: 7–10. Papazian DM, Timpe LC, Jan YN, Jan LY, 1991. Alteration of voltage-dependence of Shaker potassium channel by mutations in the S4 sequence. Nature 349: 305–310. Park KS, Yang JW, Seikel E, Trimmer JS, 2008. Potassium channel phosphorylation in excitable cells: providing dynamic functional variability to a diverse family of ion channels. Physiology (Bethesda) 23: 49–57. Persson AS, Klement G, Almgren M, Sahlholm K, Nilsson J, Petersson S, Arhem P, Schalling M, Lavebratt C, 2005. A truncated Kv1.1 protein in the brain of the megencephaly mouse: expression and interaction. BMC Neurosci 6: 65. Persson AS, Westman E, Wang FH, Khan FH, Spenger C, Lavebratt C, 2007. Kv1.1 null mice have enlarged hippocampus and ventral cortex. BMC Neurosci 8: 10. Petersson S, Persson AS, Johansen JE, Ingvar M, Nilsson J, Klement G, Arhem P, Schalling M, Lavebratt C, 2003. Truncation of the Shaker-like voltage-gated potassium channel, Kv1.1, causes megencephaly. Eur J Neurosci 18: 3231–3240. Pongs O, 1999. Voltage-gated potassium channels: from hyperexcitability to excitement. FEBS Lett 452: 31–35. Pongs O, Leicher T, Berger M, Roeper J, Bahring R, Wray D, Giese KP, Silva AJ, Storm JF, 1999. Functional and molecular aspects of voltage-gated Kþ channel beta subunits. Ann N Y Acad Sci 868: 344–355. Rasband MN, Trimmer JS, 2001. Developmental clustering of ion channels at and near the node of Ranvier. Dev Biol 236: 5–16. Rasband MN, Trimmer JS, Schwarz TL, Levinson SR, Ellisman MH, Schachner M, Shrager P, 1998. Potassium channel distribution, clustering, and function in remyelinating rat axons. J Neurosci 18: 36–47. Rasband MN, Trimmer JS, Peles E, Levinson SR, Shrager P, 1999. Kþ channel distribution and clusteringindevelopingandhypomyelinatedaxonsoftheopticnerve.JNeurocytol28:319–331. Rasband MN, Park EW, Zhen D, Arbuckle MI, Poliak S, Peles E, Grant SG, Trimmer JS, 2002. Clustering of neuronal potassium channels is independent of their interaction with PSD-95. J Cell Biol 159: 663–672.

REFERENCES

151

Rasband MN, 2004. It’s “juxta” potassium channel! J Neurosci Res 76: 749–757. Rettig J, Heinemann SH, Wunder F, Lorra C, Parcej DN, Dolly JO, Pongs O, 1994. Inactivation properties of voltage-gated Kþ channels altered by presence of beta-subunit. Nature 369: 289–294. Rhodes KJ, Keilbaugh SA, Barrezueta NX, Lopez KL, Trimmer JS, 1995. Association and colocalization of Kþ channel alpha- and beta-subunit polypeptides in rat brain. J Neurosci 15: 5360–5371. Rhodes KJ, Monaghan MM, Barrezueta NX, Nawoschik S, Bekele-Arcuri Z, Matos MF, Nakahira K, Schechter LE, Trimmer JS, 1996. Voltage-gated Kþ channel beta subunits: expression and distribution of Kv beta 1 and Kv beta 2 in adult rat brain. J Neurosci 16: 4846–4860. Rhodes KJ, Strassle BW, Monaghan MM, Bekele-Arcuri Z, Matos MF, Trimmer JS, 1997. Association and colocalization of the Kvbeta1 and Kvbeta2 beta-subunits with Kv1 alphasubunits in mammalian brain Kþ channel complexes. J Neurosci 17: 8246–8258. Rivera J, Chu PJ, Lewis TL Jr, Arnold DB, 2007. The role of Kif5B in axonal localization of Kv1 Kþ channels. Eur J Neurosci 25: 136–146. Salkoff L, Baker K, Butler A, Covarrubias M, Pak MD, Wei A, 1992. An essential “set” of Kþ channels conserved in flies, mice and humans. Trends Neurosci 15: 161–166. Santacruz-Toloza L, Huang Y, John SA, Papazian DM, 1994. Glycosylation of shaker potassium channel protein in insect cell culture and in Xenopus oocytes. Biochemistry 33: 5607–5613. Scannevin RH, et al. 2004. Two N-terminal domains of Kv4 Kþ channels regulate binding to and modulation by KChIP1. Neuron 41: 587–598. Schulte U, Thumfart JO, Klocker N, Sailer CA, Bildl W, Biniossek M, Dehn D, Deller T, Eble S, Abbass K, Wangler T, Knaus HG, Fakler B, 2006. The epilepsy-linked Lgi1 protein assembles into presynaptic Kv1 channels and inhibits inactivation by Kvbeta1. Neuron 49: 697–706. Scott VE, Muniz ZM, Sewing S, Lichtinghagen R, Parcej DN, Pongs O, Dolly JO, 1994a. Antibodies specific for distinct Kv subunits unveil a heterooligomeric basis for subtypes of alpha-dendrotoxin-sensitive Kþ channels in bovine brain. Biochem 33: 1617–1623. Scott VE, Rettig J, Parcej DN, Keen JN, Findlay JB, Pongs O, Dolly JO, 1994b. Primary structure of a beta subunit of alpha-dendrotoxin-sensitive Kþ channels from bovine brain. Proc Natl Acad Sci USA 91: 1637–1641. Sewing S, Roeper J, Pongs O, 1996. Kv beta 1 subunit binding specific for shaker-related potassium channel alpha subunits. Neuron 16: 455–463. Shamotienko OG, Parcej DN, Dolly JO, 1997. Subunit combinations defined for Kþ channel Kv1 subtypes in synaptic membranes from bovine brain. Biochemistry 36: 8195–8201. Sheng M, Sala C, 2001. PDZ domains and the organization of supramolecular complexes. Annu Rev Neurosci 24: 1–29. Sheng M, Tsaur ML, Jan YN, Jan LY, 1992. Subcellular segregation of two A-type Kþ channel proteins in rat central neurons. Neuron 9: 271–284. Sheng M, Liao YJ, Jan YN, Jan LY, 1993. Presynaptic A-current based on heteromultimeric Kþ channels detected in vivo. Nature 365: 72–75. Sheng M, Tsaur ML, Jan YN, Jan LY, 1994. Contrasting subcellular localization of the Kv1.2 Kþ channel subunit in different neurons of rat brain. J Neurosci 14: 2408–2417.

152

SHAKER FAMILY KV1 VOLTAGE-GATED POTASSIUM CHANNELS

Shi G, Trimmer JS, 1999. Differential asparagine-linked glycosylation of voltage-gated Kþ channels in mammalian brain and in transfected cells. J Membr Biol 168: 265–273. Shi G, Nakahira K, Hammond S, Rhodes KJ, Schechter LE, Trimmer JS, 1996. Beta subunits promote Kþ channel surface expression through effects early in biosynthesis. Neuron 16: 843–852. Shibata R, Misonou H, Campomanes CR, Anderson AE, Schrader LA, Doliveira LC, Carroll KI, Sweatt JD, Rhodes KJ, Trimmer JS, 2003. A fundamental role for KChIPs in determining the molecular properties and trafficking of Kv4.2 potassium channels. J Biol Chem 36445–36454. Shu Y, Yu Y, Yang J, McCormick DA, 2007. Selective control of cortical axonal spikes by a slowly inactivating Kþ current. Proc Natl Acad Sci USA 104: 11453–11458. Smart SL, Lopantsev V, Zhang CL, Robbins CA, Wang H, Chiu SY, Schwartzkroin PA, Messing A, Tempel BL, 1998. Deletion of the K(V)1.1 potassium channel causes epilepsy in mice. Neuron. 20: 809–819. Sokolova O, Accardi A, Gutierrez D, Lau A, Rigney M, Grigorieff N, 2003. Conformational changes in the C terminus of Shaker Kþ channel bound to the rat Kvbeta2-subunit. Proc Natl Acad Sci USA 100: 12607–12612. Soler-Llavina GJ, Chang TH, Swartz KJ, 2006. Functional interactions at the interface between voltage-sensing and pore domains in the Shaker K(v) channel. Neuron 52: 623–634. Storm JF, 2000. Kþ channels and their distribution in large cortical pyramidal neurones. J Physiol 525: 565–566. Stuhmer W, Ruppersberg JP, Schroter KH, Sakmann B, Stocker M, Giese KP, Perschke A, Baumann A, Pongs O, 1989. Molecular basis of functional diversity of voltage-gated potassium channels in mammalian brain. EMBO J 8: 3235–3244. Swartz KJ, 2004. Towards a structural view of gating in potassium channels. Nat Rev Neurosci 5: 905–916. Tang BL, 2001. Protein trafficking mechanisms associated with neurite outgrowth and polarized sorting in neurons. J Neurochem 79: 923–930. Tempel BL, Papazian DM, Schwarz TL, Jan YN, Jan LY, 1987. Sequence of a probable potassium channel component encoded at Shaker locus of Drosophila. Science 237: 770–775. Thornhill WB, Wu MB, Jiang X, Wu X, Morgan PT, Margiotta JF, 1996. Expression of Kv1.1 delayed rectifier potassium channels in Lec mutant Chinese hamster ovary cell lines reveals a role for sialidation in channel function. J Biol Chem 271: 19093–19098. Tiffany AM, Manganas LN, Kim E, Hsueh YP, Sheng M, Trimmer JS, 2000. PSD-95 and SAP97 exhibit distinct mechanisms for regulating Kþ channel surface expression and clustering. J Cell Biol 148: 147–158. Tipparaju SM, Liu SQ, Barski OA, Bhatnagar A, 2007. NADPH binding to beta-subunit regulates inactivation of voltage-gated Kþ channels. Biochem Biophys Res Commun 359: 269–276. Trimmer JS, 1998. Analysis of Kþ channel biosynthesis and assembly in transfected mammalian cells. Methods Enzymol 293: 32–49. Trimmer JS, Rhodes KJ, 2004. Localization of voltage-gated ion channels in mammalian brain. Annu Rev Physiol 66: 477–519. Tsai W, Morielli AD, Peralta EG, 1997. The m1 muscarinic acetylcholine receptor transactivates the EGF receptor to modulate ion channel activity. EMBO J 16: 4597–4605.

REFERENCES

153

Tytgat J, Debont T, Carmeliet E, Daenens P, 1995. The alpha-dendrotoxin footprint on a mammalian potassium channel. J Biol Chem 270: 24776–24781. Vabnick I, Trimmer JS, Schwarz TL, Levinson SR, Risal D, Shrager P, 1999. Dynamic potassium channel distributions during axonal development prevent aberrant firing patterns. J Neurosci 19: 747–758. Vacher H, Misonou H, Trimmer JS, 2006. Determinants of voltage-gated potassium channel distribution in mammalian neurons. In: Bean AJ (Ed.), Protein Trafficking in Neurons. Elsevier Academic Press, London, pp. 244–270. Vacher H, Mohapatra DP, Misonou H, Trimmer JS, 2007. Regulation of Kv1 channel trafficking by the mamba snake neurotoxin dendrotoxin K. FASEB J 21: 906–914. Van Wart A, Trimmer JS, Matthews G, 2007. Polarized distribution of ion channels within microdomains of the axon initial segment. J Comp Neurol 500: 339–352. Veh RW, Lichtinghagen R, Sewing S, Wunder F, Grumbach IM, Pongs O, 1995. Immunohistochemical localization of five members of the Kv1 channel subunits: contrasting subcellular locations and neuron-specific co- localizations in rat brain. Eur J Neurosci 7: 2189– 2205. Wang H, Kunkel DD, Martin TM, Schwartzkroin PA, Tempel BL, 1993. Heteromultimeric Kþ channels in terminal and juxtaparanodal regions of neurons. Nature 365: 75–79. Wang H, Kunkel DD, Schwartzkroin PA, Tempel BL, 1994. Localization of Kv1.1 and Kv1.2, two K channel proteins, to synaptic terminals, somata, and dendrites in the mouse brain. J Neurosci 14: 4588–4599. Watanabe I, Wang HG, Sutachan JJ, Zhu J, Recio-Pinto E, Thornhill WB, 2003. Glycosylation affects rat Kv1.1 potassium channel gating by a combined surface potential and cooperative subunit interaction mechanism. J Physiol 550: 51–66. Weng J, Cao Y, Moss N, Zhou M, 2006. Modulation of voltage-dependent Shaker family potassium channels by an aldo-keto reductase. J Biol Chem 281: 15194–15200. Wenzel HJ, Vacher H, Clark E, Trimmer JS, Lee AL, Sapolsky RM, Tempel BL, Schwartzkroin PA, 2007. Structural consequences of Kcna1 gene deletion and transfer in the mouse hippocampus. Epilepsia 48: 2023–2046. Winklhofer M, Matthias K, Seifert G, Stocker M, Sewing S, Herget T, Steinhauser C, SaalerReinhardt S, 2003. Analysis of phosphorylation-dependent modulation of Kv1.1 potassium channels. Neuropharmacology 44: 829–842. Yang JW, Vacher H, Park KS, Clark E, Trimmer JS, 2007. Trafficking-dependent phosphorylation of Kv1. 2 regulates voltage-gated potassium channel cell surface expression. Proc Natl Acad Sci USA 104: 20055–20060. Yellen G, 1998. The moving parts of voltage-gated ion channels. Q Rev Biophys 31: 239–295. Yu W, Xu J, Li M, 1996. NAB domain is essential for the subunit assembly of both alpha-alpha and alpha-beta complexes of shaker-like potassium channels. Neuron 16: 441–453. Yuan LL, Chen X, 2006. Diversity of potassium channels in neuronal dendrites. Prog Neurobiol 78: 374–389. Zerr P, Adelman JP, Maylie J, 1998. Episodic ataxia mutations in Kv1. 1 alter potassium channel function by dominant negative effects or haploinsufficiency. J Neurosci 18: 2842–2848. Zhu J, Watanabe I, Gomez B, Thornhill WB, 2001. Determinants involved in Kv1 potassium channel folding in the endoplasmic reticulum, glycosylation in the Golgi, and cell surface expression. J Biol Chem 276: 39419–39427.

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Zhu J, Watanabe I, Gomez B, Thornhill WB, 2003. Heteromeric Kv1 potassium channel expression: amino acid determinants involved in processing and trafficking to the cell surface. J Biol Chem 278: 25558–25567. Zhu J, Gomez B, Watanabe I, Thornhill WB, 2005. Amino acids in the pore region of Kv1 potassium channels dictate cell-surface protein levels: a possible trafficking code in the Kv1 subfamily. Biochem J 388: 355–362.

6 UNIQUE MITOCHONDRIAL ION CHANNELS: ROLES IN SYNAPTIC TRANSMISSION AND PROGRAMMED CELL DEATH Elizabeth A. Jonas Department of Internal Medicine, Yale University, New Haven, CT 06520, USA

6.1 INTRODUCTION Mitochondria produce ATP for a myriad of cellular processes including the highly energy-dependent events of synaptic transmission. Mitochondria exist at the center of a web of signaling within the cell that regulates cell fate. They contribute to a spectrum of cell death types from necrosis to apoptosis. In many forms of programmed cell death, mitochondria play a crucial role in regulating oxidative phosphorylation to produce or conserve precious energy supplies. They also manage cytosolic levels of calcium and zinc, key ions implicated in excitotoxic neuronal death. This type of death has been implicated in cellular metabolic decline, neurodegeneration, and aging. Mitochondria contain two membranes, an inner and an outer membrane. In canonical apoptotic death, mitochondria regulate the release of proapoptotic factors such as cytochrome c from the mitochondrial intermembrane space across the outer membrane. Cytochrome c and other proapoptotic molecules activate downstream enzymes, including caspases that destroy the cell contents and its nuclear material. Outer membrane mitochondrial voltage-gated ion channels are intimately involved in these cellular processes. These ion channels regulate the Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

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release of ATP and the reuptake of ADP through mitochondrial membranes, the influx and efflux of calcium, sodium, potassium, and zinc, which help determine the membrane potential of the inner membrane and regulate matrix volume. In addition, a family of proteins known as the BCL-2 family, which reside in the outer membrane and inhibit the release of proapoptotic factors from mitochondria, have properties of ion channels. Knowledge of their roles in the regulation of cellular functions has been expanding. In particular, we now know that BCL-2 family proteins perform physiological functions as well as participate in cell death, including, for example, regulation of the strength and pattern of synaptic transmission. Mitochondria also use channels to import proteins from the cytosol. These channels may themselves be regulated by changes in mitochondrial membrane potential. Although little is known about how the outer membrane voltage is determined, it has become clear that its permeability is tightly controlled, in part by the voltage dependence of outer membrane conductances. The control of outer membrane permeability compartmentalizes the intermembrane space with implications for metabolic regulation and cell fate determination. Until late in the last century, investigators mainly concentrated on understanding the inner mitochondrial membrane and matrix mitochondria efficiently produce ATP by oxidative phosphorylation. Studies led to a consensus that the energy produced by the reduction of NAD(P) to NAD(P)H and FAD to FADH in the tricarboxylic acid cycle was used (by reoxidizing these species in the oxidation part of the reaction) for the transport of electrons from high to low energy electron acceptors and to pump Hþ ions out across the inner membrane, thereby producing a large electrochemical (voltage) gradient. The movement of Hþ ions down the gradient across the ATP synthase phosphorylates ADP, producing ATP (the phosphorylation part of the reaction). The inner mitochondrial membrane is a barrier similar to the plasma membrane that allows the separation of ion species to produce the electrochemical gradient. The potential energy produced by charge separation across the inner membrane can be used to produce work and for signaling through the actions of its ion channels. These channels regulate the ATP-producing activity of the mitochondria and the enzymatic activity of the matrix. They control cytosolic concentrations of several ions and they alter the concentration of signaling molecules. The chapter is divided into three parts, addressing the unique aspects of mitochondrial ion channel function in the nervous system. The first part will summarize the current knowledge of the functions of the unique mitochondrial channels in the outer membrane, including the voltage-dependent anion channel (VDAC) and the channels of the BCL-2 family. The second part will address the known unique mitochondrial inner membrane ion channels that regulate ion flux into and out of the matrix and will suggest that the outer and inner membrane channels exist in a protein complex that is highly regulated. The third part will address the combined the functions of the inner and outer membrane channels, including those of the BCL-2 family, and discuss how regulation of such channels may alter the physiological behavior of the neuronal synapse.

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6.2 OUTER MITOCHONDRIAL MEMBRANE CHANNELS 6.2.1

Role of VDAC in Mitochondrial Function

Perhaps the most prevalent ion channel in the mitochondrial outer membrane is the voltage-dependent anion channel (mitochondrial porin, VDAC), and no studies of mitochondria, metabolism, or cell death can ignore the biophysical characteristics or physiological behavior of this important molecule (Colombini et al., 1996; ShoshanBarmatz et al., 2006). The main function of VDAC appears to be its ability to conduct metabolites such as ATP, ADP, NADH, and pyruvate, in addition to metabolites whose molecular weight can reach almost 1000. VDAC, therefore, has different biophysical characteristics than other voltage-gated ion channels (Hodge and Colombini, 1997). Historically, it was thought that VDAC was always open, making the mitochondrial outer membrane like a leaky sieve, but many recent studies have contributed to the present notion that the opening and closing of VDAC are highly regulated (Kinnally and Tedeschi, 1994; Jonas et al., 1999). The 30 kDa VDAC protein is relatively small compared to the plasma membrane voltage-gated ion channels such as Naþ or Caþ channels, whose molecular weights are approximately 300 kDa. VDAC is highly conserved in tertiary structure from yeast to man and it is also clearly present in plants. Interestingly, different VDACs differ widely in their amino acid sequences even though their channel gating and selectivity properties are conserved, suggesting that different interacting partners may bind to alternative sequences to regulate channel behavior (Colombini et al., 1996). 6.2.1.1 Biophysical Characteristics of VDAC In all the VDAC sequences analyzed, the N terminus contains a sequence that can form an amphiphilic alpha helix. All the downstream sequences have 12 or more alternating hydrophobic/ hydrophilic segments, which form the beta sheet that makes up the wall of a cylinder or barrel (Blachly-Dyson et al., 1990). In this configuration, the hydrophilic inner surface through which water and solutes travel faces the pore and the hydrophobic outer surface is buried in the membrane (Blachly-Dyson et al., 1989). Site-directed mutagenesis confirmed that residues in the alpha helix and the beta sheet strands are necessary to define selectivity, whereas some residues that do not affect selectivity are assigned positions outside the transmembrane region (Colombini et al., 1996). All VDACs studied have a conserved set of biophysical features (Colombini, 1989; Colombini et al., 1996). The channels have a very large single-channel conductance of 1–4 nS when the purified protein is reconstituted into artificial lipid membranes. The open state of the channel prefers anions to cations 2:1 and also strongly prefers metabolically relevant anions such as ATP. The channels open at voltages close to 0 mV, either in the positive or negative direction, but undergo rapid transitions to several closed states at potentials less than
50 or more than þ50 (Fig. 6.1). These “closed” states are permeable to cations but not to anions such as metabolites. Some closed states are more stable than others, so that the channel tends to reopen from some of the states more frequently than from others. Since the selectivity of the channel

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Membrane conductance (pS)

(a)

Selectivity filter

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4 Amount pos. charge in pore: low Amount pos. charge in pore: high 3 2

+ + +

1 0 -100 -80 -60 -40 -20 0

20 40 60 80 100

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Gating

“Closed state”-cation permeable

FIGURE 6.1 Voltage dependence of membrane conductance of the voltage-dependent anion channel (VDAC). (a) The artist’s rendition simulates data recorded when the channel protein is reconstituted in artificial membranes. Increasing the number of charged residues in the pore increases the steepness of the voltage dependence. (b) Dependence of the open and closed states of VDAC on position of the charged selectivity moiety inside or outside of the pore.

changes dramatically upon gating, the electrostatic forces within the ion conducting pathway must also change. Certain residues therefore affect the channel behavior in the open state but not in the closed state. This finding suggests that a positively charged moiety moves out of the pore during gating (Peng et al., 1992) (Fig. 6.1). The removal of positive charges may lead to a decrease in the propensity to conduct anions (such as metabolites) in the closed state. The change should also result in a reduction in the volume of water in the pore and a reduction in the pore radius. The change in selectivity with gating also predicts that the positive charges that move during gating may comprise the voltage sensor (Colombini et al., 1996). Therefore, increasing the positive charge within the pore will increase the steepness of voltage dependence and decreasing the positive charge will decrease the steepness (Fig. 6.1) (Doring and Colombini, 1985). Interestingly, moving the sensor against a high salt gradient requires more energy than moving it down the salt gradient; therefore, it appears that kinetic energy from the movement of the salt may help move the sensor out of the channel (Zizi et al., 1998). Regulators of VDAC include a variety of polyanions, all of which have similar effects (Colombini, 1989). When they are added to one side of the membrane, they appear to favor the closure of the channel, possibly by drawing out the positively charged gate region (Mangan and Colombini, 1987). Konig’s polyanion (a VDAC inhibitor that is a copolymer of styrene, maleic acid, and methacrylic acid) increases the probability of the channel closing even in the absence of a membrane potential, suggesting that its hydrophobic regions allow it to bind tightly to the membrane as well as to the voltage sensor (Colombini et al., 1987). It induces a progressive decrease in conductance of the channel over time as the channel enters several closed states. In addition to exogenously applied inhibitors, a large (100,000 MW) endogenous soluble protein modulator of VDAC, most likely residing within the intermembrane space,

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also increases the voltage-dependence of the channel and favors the closed state (Holden and Colombini, 1988). Of the nucleotides that penetrate the open channel, only NADH and NADPH significantly affect channel gating. NADH doubles the voltage-dependence of VDAC (Zizi et al., 1994). The addition of NADH and NADPH to mitochondria with intact outer membranes results in a six-fold reduction of the permeability of the outer membrane to ADP (Lee et al., 1994). VDAC may favor the closed state under normal conditions due to the presence of negative anions and protein modulators in the intermembrane space. Immobile molecules in the intermembrane space may exert osmotic pressure, drawing water through the pore and decreasing hydrostatic pressure within the pore (Holden and Colombini, 1993). The decline in hydrostatic pressure decreases pore volume and finally favors channel closure (Zimmerberg and Parsegian, 1986). 6.2.1.2 Control of Metabolism by VDAC Hexokinase is the first enzyme in the glycolytic pathway (Lemasters and Holmuhamedov, 2006) and it requires ATP. It apparently binds to the outer surface of the outer membrane via its association with VDAC, giving it preferential access to the ATP that is leaving the mitochondrion and also giving the mitochondrion preferential access to ADP produced by hexokinase. Hexokinase thereby uses ATP that is made in mitochondria over exogenous ATP. The relationship between VDAC and hexokinase suggests that VDAC is the central regulator of the interaction between glycolysis and mitochondrial respiration (Golshani-Hebroni and Bessman, 1997). Contact sites between the two membranes may contain the colocalized adenine nucleotide transporter (ANT, transporter of ADP and ATP through the inner membrane), VDAC, and hexokinase (Hashimoto and Wilson, 2000). Increasing the proximity of VDAC to the inner membrane proteins provides ADP readily to the ANT, thereby possibly increasing the efficiency of

FIGURE 6.2 Regulation of mitochondrial metabolism by interaction of hexokinase with VDAC. On the left is shown the case where hexokinase is not bound to VDAC. Inefficient transfer of ATP and ADP across the two membranes leads to the loss of crosstalk between glycolysis and mitochondrial respiration in the matrix. On the right is shown the case where hexokinase is bound to VDAC. Efficient exchange of metabolites occurs, increasing crosstalk between glycolysis and mitochondrial respiration.

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production of ATP (Fig. 6.2). An increase in metabolic rate, by activating glycolysis and hexokinase, increases the likelihood that the two membranes will come into close contact (Hackenbrock, 1972a; Brdiczka et al., 1986). The ATP generated by mitochondrial respiration then further stimulates the enzymatic activity of hexokinase. In certain tumor cells, hexokinase is very highly expressed. In these cells, the unusually high amount of hexokinase may cause VDAC to move closer to the inner membrane, increasing the association between glycolysis and mitochondrial metabolism (GolshaniHebroni and Bessman, 1997). Some tumor cells may lack this type of modulation. In these cells, glucose inhibits rather than activates oxidative phosphorylation in a reaction known as the Crabtree effect. High hexokinase levels cause hexokinase to bind tightly to VDAC, inhibiting the conductance of metabolites (Colombini et al., 1996; Penso and Beitner, 1998). In these defective cells, the ANT is not present at the hexokinase sites and contact sites are absent (Denis-Pouxviel et al., 1987). However, the cells can be converted to normal metabolism by lowering glucose levels, thereby enhancing mitochondrial metabolism, which causes contact sites and VDAC/ANT connections to re-form. Hexokinase, therefore, can help the cell switch between glycolysis and mitochondrial metabolism. When hexokinase inhibits VDAC, presumably no mitochondrial ATP can exit the mitochondrion nor can ADP enter; therefore, the cell must switch from mitochondrial metabolism to glycolysis. However, the homeostatic mechanisms regulating VDAC closure are quite complex since glucose-6-phosphate, the product of the enzymatic activity of hexokinase, antagonizes the binding of hexokinase to VDAC and tends to favor the open state. Therefore, if glycolysis becomes inhibited downstream of glucose-6-phosphate, glucose-6-phosphate will accumulate and then VDAC may open again. 6.2.1.3 VDAC Structural Studies EM studies suggest that monomeric VDAC channels form a crystalline array on the outer membrane (Rostovtseva et al., 2005). A pore of 2.5–3 nm was suggested by EM studies and confirmed by the inability of nonelectrolytes to penetrate through the narrowest portion of the pore (Rostovtseva et al., 2005). When the channel is in its closed state, a rigid spheroid molecule of 1.9 nm (gamma cyclodextrin) can barely permeate. Studies of these nonpermeating macromolecules, which favor closure by producing tension within the channel, show that the change in channel volume upon closure is 20–40 nm3. This result supports a model whereby a global change in the pore structure occurs, rather than a local constriction at one point within the pore. 6.2.2

BCL-2 Family Ion Channels: Role in Programmed Cell Death in Neurons

Programmed cell death or apoptosis is the genetic predisposition of cells to die (Adams and Cory, 2007). In this fashion, cells are eliminated developmentally and throughout the life of an organism to remove old and damaged cells (Kroemer and Reed, 2000). Failure of the death program can lead to the unchecked growth of cancer cells, while untimely onset of cell death leads to degenerative changes. In the nervous system, premature cell death produces diseases such as Alzheimer’s disease and amyotrophic lateral sclerosis (Yuan and Yankner, 2000). In addition, during pathological brain insults such as ischemia, infection, or trauma, some brain cells die immediately while

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others die a delayed death long after the insult, by turning on programmed death pathways (Banasiak et al., 2000). A highly evolutionarily conserved form of cell death in vertebrate cells is organized by mitochondria, which appear to receive a signal to undergo permeabilization of their outer membranes (Green and Kroemer, 2004; Adams and Cory, 2007). The opening of the outer mitochondrial membrane is also sometimes referred to as the activation of the apoptosis channel (Dejean et al., 2005). This event occurs suddenly, leading to the release of several intermembrane space proteins such as cytochrome c (Green and Kroemer, 2004; Martinez-Caballero et al., 2005). The release of cytochrome c produces two events. On the one hand, the loss of cytochrome c compromises the ability of mitochondria to produce ATP and eventually to maintain the mitochondrial membrane potential. On the other hand, cytochrome c and the other factors released from the mitochondria activate downstream caspases that chew up hundreds of cellular proteins (Youle and Strasser, 2008). Exactly how mitochondrial membrane permeabilization occurs is still unclear. Nevertheless, it appears that the BCL-2 family proteins play a major role in either regulating or, through their ion channel function, by producing permeabilization of the outer membrane (Fig. 6.3). BCL-2 family proteins were first described when the gene encoding BCL-2 was identified in B-cell lymphomas (Tsujimoto et al., 1985). BCL-2 family proteins regulate the protection of cells against prodeath signals, including growth factor

FIGURE 6.3 Interaction of inner and outer mitochondrial membrane ion channels in different scenarios. (From left to right): (1) The release of cytochrome c from the intermembrane space during cell death through an outer membrane channel formed by Bax or a VDAC/D N BCL-xL complex. (2) During physiological functioning of mitochondria such as during cell growth or during inhibition of cell death, VDAC may open to release metabolites, assisted by interaction with BCL-xL. (3) Formation of a two-membrane-spanning complex of channel proteins. If the inner membrane channel is activated by the binding of cyclophilin D in the presence of calcium, an outer membrane channel such as VDAC may open in response to activation of a messenger within the intermembrane space, or (4) by formation of a channel complex spanning the two membranes. If the mPTP is activated, swelling of the mitochondrial matrix followed by bursting of the outer membrane may cause release of cytochrome c from the intermembrane space.

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deprivation, ultraviolet and gamma radiation, heat shock, tumor necrosis factor, viral infections, and free radical formation (Adams and Cory, 2007). In addition, BCL-2 family proteins regulate death stimuli that had previously been relegated to the realm of necrosis, such as ischemic cell death in the heart and brain (Bonanni et al., 2006). 6.2.2.1 BCL-2 Family Proteins Function as Ion Channels BCL-2 family proteins exhibit two important features: They produce ion channel activity in intracellular organelle membranes, and they are regulated by interactions with binding partners, including BCL-2 family members and other intracellular membrane components. As a result, the BCL-2 family proteins control the release of death-inducing mitochondrial factors such as cytochrome c, which is normally present in the intermembrane space, into the cytosol (Fig. 6.3). Certain BCL-2 family proteins may also enhance the release of metabolites from mitochondria by interaction with VDAC (Fig. 6.3) (Gottlieb et al., 2002) and interact with the members of the growth and survival signaling pathway (Zha et al., 1996). In some cases, antiapoptotic proteins can be rapidly converted into proapoptotic molecules by proteolytic cleavage after a cell death stimulus, such as the transformation of BCL-xL to DN BCL-xL, and this event dramatically increases the permeability of the mitochondrial outer membrane by increasing the single-channel conductance of the pore-forming protein (Fig. 6.3) (Cheng et al., 1997; Clem et al., 1998; Jonas et al., 2004). In neurons, the size of the mitochondrial channel conductance and its binding partners may determine whether a mitochondrial channel promotes cell survival or death. Both pro- and antiapoptotic BCL-2 family proteins produce ion channel activity when inserted into artificial lipid membranes (Schlesinger et al., 1997; Schendel et al., 1998). Many BCL-2 family proteins are localized to the outer mitochondrial membrane or translocated into the outer membrane upon a death stimulus (Wolter et al., 1997; Kaufmann et al., 2003). Three categories of BCL-2 proteins contribute to the regulation of cell death (Fig. 6.4). These are the antiapoptotic members (such as

FIGURE 6.4 BCL-2 family proteins. Canonical antiapoptotic proteins contain the BH4 domain that is lacking in proapoptotic BCL-2 family members such as cleaved BCL-xL (DN BCL-xL) and Bax. BH3-only molecules are traditionally thought to be proapoptotic, but they may also play physiological roles.

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BCL-xL, BCL-2, and MCL-1), the proapoptotic members such as Bax and Bak, and a large group of BH3-only proteins such as BID, BAD, PUMA, and NOXA (Galonek and Hardwick, 2006). The antiapoptotic members of the group are similar in structure and sequence to proapoptotic Bax and Bak, and in addition to the BH1–3 domains, contain a BH4 domain that is important for the antiapoptotic features of the molecules (Tsujimoto and Shimizu, 2000; Sugioka et al., 2003). Despite the presence of the BH4 domain in BCL-xL, at first glance, the ion channel activities of Bax and BCL-xL seem quite similar (Schlesinger et al., 1997). The three-dimensional structure of BCL-xL is comprised of seven alpha helices (Muchmore et al., 1996; Schendel et al., 1998). Two outer layers of amphipathic helices serve to screen the long hydrophobic alpha helices from the aqueous domain. A long proline-rich loop found between the first and second helices is absent in the proapoptotic members of the family. The loop may be vulnerable to protease digestion and contains phosphorylation sites. The BH1–3 domains of BCL-xL fold together to give a hydrophobic region involved in homo- and heterodimerization and contain domains important for interaction with proteins that contain only a BH3 domain. The structure of BCL-xL is strikingly similar to that of the diphtheria toxin membrane translocation domain and the pore-forming domains of bacterial colicins that kill sensitive cells via the formation of a highly conductive ion channel in the target cell’s plasma membrane. The overall organization of the colicin-like channels is that of a hydrophobic region containing the pore, shielded by amphipathic helices that keep the molecule soluble in the cytoplasm until insertion into a membrane activates the pore function. Two helices of Bax and BCLxL are insufficient to form a pore (Schendel et al., 1998) but their ability to homo- and heterodimerize may provide for interactions critical for their pore-forming ability. It is possible that channel activity observed in artificial lipid membranes is not the same as activity in vivo. Both BCL-xL and Bax have demonstrated avid pore-forming capability in lipid bilayers if lipid composition of the bilayer is carefully regulated (Minn et al., 1997; Schlesinger et al., 1997; Schendel et al., 1998; Antonsson et al., 2000). The BCL-xLchannel inlipidbilayers isa nonselectivechannel that favors the conductanceof cations over anions and displays multiple conductances, with the most common conductance value of about 276 pS (Minn et al., 1997). Both Bax and BCL-xL display similar channel activity with multiple conductances, but BCL-xL has a linear conductancewhereasBaxappears to be more rectified toward positive potentials, is more anionselective than BCL-xL, and has larger peak conductances (Schlesinger et al., 1997). Channel Activity of BCL-2 Proteins Recorded in Vivo The similar channel activities in these anti- and proapoptotic proteins raised several questions, the most important of which was whether the channel activity of the anti- and proapoptotic molecules was important at all for their anti- and proapoptotic functions, and if so,why were the channel activities so similar and could they prove more dissimilar in vivo? To examine these issues, the giant synapse of the squid stellate ganglion has been used as a model system to study the activity of the recombinant proteins in mitochondrial membranes inside a living neuron (Jonas et al., 2003, 2004). In this method, a concentric electrode arrangement, a clean patch pipette tip of small internal diameter is exposed to mitochondria inside the neuronal presynaptic ending, and channel activity

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FIGURE 6.5 Channel recordings performed on mitochondrial membranes within a living neuronal synapse. (a) A control recording and recordings of patches exposed to recombinant FL BCL-xL or Bax. (b) The panel shows recordings of patches exposed to Bax or DN BCL-xL, demonstrating the large conductance activity of the membrane in the presence of these proteins. (c) A mitochondrial recording within a hypoxic synapse and a recording of a patch exposed to a VDAC inhibitor (NADH) within a hypoxic synapse.

is recorded from a patch of outer mitochondrial membrane exposed to the inside of the pipette. When such patches are exposed to recombinant antiapoptotic full-length BCL-xL protein (FL BCL-xL), mitochondrial channel activity that is distinctly different from that of controls appears (Fig. 6.5). Application of FL BCL-xL produces characteristic activity with multiple conductances in mitochondrial patches within 5 min of the start of the recording. Unitary openings correspond to conductances between 100 and 760 pS. Channel activity switches rapidly between different conductance levels. Holding the voltage at different potentials reveals a current voltage relationship that is linear or very slightly outwardly rectifying (Jonas et al., 2003). Channel activity of recombinant Bax recorded in presynaptic mitochondrial membranes shares some features with that of recombinant FL BCL-xL. To form channels readily in vitro, however, it must be activated by treatment with detergent, which causes the protein to oligomerize (Hsu and Youle, 1998; Antonsson et al., 2000; Green and Kroemer, 2004). In isolated mitochondria or isolated outer mitochondrial membranes, this activation of Bax is mediated by endogenous tBID (N-terminally cleaved/truncated BID) (Roucou et al., 2002) or another BH3-only protein (Polster et al., 2001), which may also be present in squid (Jonas et al., 2005). The findings suggest that activation of

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Bax channel activity requires a mitochondrial membrane component (Roucou et al., 2002). When purified recombinant Bax protein lacking the C terminus (BaxDC) or full-length Bax protein is placed inside the patch pipette used to record from mitochondria in the squid presynaptic terminal, it evokes channel activity in the patches ranging between 100 and 750 pS and demonstrates outward rectification (Fig. 6.5). In 5–10% of the channels observed on mitochondria inside the neurons with recombinant Bax, a number of large openings with conductances >750 pS are detected in addition to the smaller conductances, similar to the large conductances reported for purified Bax in artificial lipid membranes (Dejean et al., 2005). In contrast to the intermediate conductance openings, the current–voltage relations for the large openings are linear (Fig. 6.5). Thus, it appears that including Bax in the patch pipette induces channel activity with two distinct properties, a large conductance state and a less conductive state that shares some properties with that induced by full-length BCL-xL. The findings suggest that the smaller conductance channel activity of Bax could represent the activity of Bax in a form prior to its exposure to a death stimulus or to activator BH3 peptides. However, this “inactive” form of Bax could have additional functions besides waiting to bring on cell death in response to a stimulus. Bax may alter synaptic function in the healthy cell. The evidence that Bax and Bak can act as prosurvival factors in neurons exists (Lewis et al., 1999; Middleton and Davies, 2001; Fannjiang et al., 2003). Both in vitro and in vivo data support the hypothesis that the functional activities of Bax depend on the specific cell in which Bax resides and the specific pathological stress that the cell is undergoing. It is possible that the protective effects of Bax and Bak in such models may result indirectly from their actions on synaptic activity (Fannjiang et al., 2003; Jonas et al., 2005). If the smaller conductance activity represents the activity of Bax in healthy cells, then the infrequently detected large conductance activity could be important for its death-promoting actions. Indeed, proteolytic cleavage of Bax accelerates the onset of its prodeath function (Wood and Newcomb, 2000) and it is conceivable that the observed spontaneous large activity could have been recorded in synapses in which a death stimulus had been activated by previous damage. 6.2.2.2 Endogenous Death Channels Produced by Bax-Containing Protein Complexes The first patch-clamp recordings of endogenous death channel activity were performed on mitochondrial outer membranes isolated from cells undergoing apoptosis (Pavlov et al., 2001). Pavlov et al. were able to detect an ion channel, mitochondrial apoptosis-induced channel (MAC), whose pore diameter was estimated to be of sufficient size (4 nm) to allow the passage of cytochrome c and larger proteins. The channel displays multiple conductances, the largest of which is 2.5 nS. The channel activity is expressed in mitochondrial outer membranes, inhibited in cells overexpressing BCL-2, and is similar to the activity of pure Bax expressed in artificial lipid membranes. The timing of cytochrome c release in apoptotic cells correlates well with the onset of MAC activity and with the translocation of Bax to mitochondrial membranes, further suggesting that such channel complexes include Bax protein. Moreover, MAC activity can be immunodepleted from mitochondrial membranes treated with anti-Bax antibodies.

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Inactivator BH3 protein

Activator BH3 protein

BAD

BID

BCL BCL-xL L

B Bax activation ti ti

Cytochrome c release

Caspase activation

Cell death

FIGURE 6.6 Schematic of an example of one possible hierarchical model of the control of cell death by BCL-2 family proteins. From left to right, after a cell death signal, BAD (the inactivator BH3 protein) sequesters BCL-xL, freeing BID (the activator BH3 protein) to activate Bax. Bax oligomerizes and permeabilizes mitochondrial outer membranes, releasing cytochrome c to activate downstream caspase pathways.

6.2.2.3 Actions of BCL-2 Family Proteins are Regulated by Binding Partners The mechanisms by which antiapoptotic proteins such as BCL-2 and BCL-xL prevent cell death are poorly understood. Recent studies have focused on the ability of antiapoptotic proteins such as BCL-xL to bind to and sequester proapoptotic members of the BCL-2 family (Kim et al., 2006). BCL-2 and BCL-xL bind to Bax and to BH3 peptides, preventing the proapoptotic actions of these proteins (Galonek and Hardwick, 2006; Kim et al., 2006). Under resting conditions, BCL-xL and BCL-2 are bound to activator BH3 molecules, keeping them in a quiescent state. Data suggest that activation of inactivator BH3-only proteins such as BAD is the first step in the initiation of cell death. These inactivator BH3 proteins then bind to the antiapoptotic BCL-2 proteins BCL-xL and BCL-2, preventing them from binding to Bax or to activator BH3 peptides such as BID. The activator BH3 peptides can then bind to and activate Bax or bind to inhibitors of Bak, such as VDAC2 (Cheng et al., 2003). The end result is that Bax or Bak are free to homo-oligomerize and thereby permeabilize outer mitochondrial membranes. This hierarchy model explains the high killing potency of a subset of BH3-only proteins that is known to bind all antiapoptotic proteins (Fig. 6.6) (Galonek and Hardwick, 2006; Kim et al., 2006). Bax and BCL-xL share high structural homology in the BH1–BH3 domains (Schlesinger et al., 1997; Schendel et al., 1998). It is clear from the binding pair studies that both Bax and BCL-xL interact in complexes with the BH3-only proteins. When other proteins or binding partners compete for their binding, they both may get released from these complexes and can perform other functions. 6.2.3

Interaction of VDAC with BCL-2 Family Proteins

Because of the importance of VDAC in mitochondrial function, a role for VDAC in cell death has been postulated. The possibility that VDAC is a component of the permeability transition pore, which can in the absence of other outer membrane

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proteins release cytochrome c during programmed cell death, remains controversial. Against the idea that it is the sole member of the apoptotic channel is the finding that programmed cell death can occur in the absence of VDAC (Baines et al., 2007) and that the functional unit of VDAC is most likely a monomer (Rostovtseva et al., 2005) whose pore size is too small to release proapoptotic factors such as cytochrome c. However, several reports support the tendency of VDAC to self-assemble into dimers, trimers, and tetramers (Zalk et al., 2005; Malia and Wagner, 2007).Whether these structures could actually form in mitochondrial outer membranes during cell death is not known. VDAC dimerization is produced by As2O3 (Zheng et al., 2004) during the process of release of cytochrome c, and it has been suggested that VDAC oligomerization could be dependent on the presence of cytochrome c (Zalk et al., 2005). A further argument against the release of cytochrome c by VDAC is that VDAC only passes certain negatively charged metabolites, most not much larger than ATP. Large, positively charged cytochrome c is unlikely to pass through VDAC since permeation by cations is reduced in the open conformation. VDAC may interact with molecular regulators of apoptosis and metabolism such as hexokinase on the cytosolic side and creatine kinase in the intermembrane space. The precise mechanism by which association with these molecules regulates the permeability of VDAC is unknown (Wicker et al., 1993; Brdiczka et al., 1994). A description of the studies in which VDAC has been found to interact with BCL-2 family proteins follows. 6.2.3.1 Models of VDAC -BCL-2 Family Interaction Although the diameter of VDAC at its largest is not compatible with the release of cytochrome c, it is possible that VDAC-interacting proteins could regulate its pore diameter or interact in multimeric structures with VDAC, forming a pore large enough to release proapoptotic factors from the mitochondrion. One controversial model proposes that Bax interacts with VDAC to provide a large pore through which cytochrome c could permeate (Shimizu et al., 1999). Studies by this group suggest that the single-channel conductance of Bax, when combined with VDAC in planar lipid bilayers, is increased by a factor of 4–10 over VDAC and Bax channels alone, and the group also finds that liposomes containing these proteins are permeable to cytochrome c (Shimizu et al., 2000). The authors also found that the antiapoptotic BH4 domain of BCL-2 closed VDAC and inhibits cell death (Shimizu et al., 2000). An equally controversial model suggests that BCL-2 family proteins regulate mitochondrial membrane permeability by closing VDAC during apoptosis. This closure leads to a lack of metabolite exchange across the outer membrane eventually accompanied by inner membrane swelling and rupture of the outer membrane, thereby releasing cytochrome c (Vander Heiden et al., 2000). This process is prevented by the antiapoptotic protein BCL-xL, which by interacting with VDAC after a cell death stimulus maintains metabolic exchange across the outer mitochondrial membrane and prevents cell death (Gottlieb et al., 2002). Proapoptotic tBID has also been proposed as a VDAC interacting partner. In a study of VDAC reconstituted into artificial lipid membranes, the activated proapoptotic protein BID induced VDAC channel closure (Rostovtseva et al., 2004).

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6.2.3.2 Interactions of VDAC with BCL-xL A recent structural study of BCL-xL and VDAC illuminated potential sites of interaction between the two molecules in an artificial lipid environment. Recombinantly expressed VDAC (Malia and Wagner, 2007) was dissolved in a detergent buffer that mimicked the state of VDAC within the outer mitochondrial membrane. The structural spectrum resulting from the analysis of this solution reveals a folded protein of high beta sheet content, as described previously. Titration of VDAC with a large excess of metabolites results in chemical shifts indicative of low affinity binding of the metabolites to VDAC (especially NADH, NAD, and ATP), a result predicted by the requirement of low affinity binding for efficient transport. Exposing the dissolved VDAC solution to BCL-xL also causes a marked shift of the chemical spectrum of VDAC, indicating significant interaction between VDAC and BCL-xL. In complementary experiments, VDAC in turn causes a marked shift in the spectrum of BCL-xL. These results, and those of additional studies, demonstrate that VDAC appears to bind to BCL-xL in two areas. The first is the helical hairpin region of BCL-xL near the C-terminus that is the putative site of interaction of BCL-xL with membranes. The other is a region near the N-terminus that may form part of the BH4 domain. Previous studies had suggested that the BH4 domain of BCL-xL was necessary for the inhibition of VDAC channel opening by BCL-xL (Shimizu et al., 2000). Thus these structural findings support the findings of the artificial lipid bilayer studies. The model based on these findings predicts that the two membrane proteins are oriented parallel to each other in the membrane and bind to each other along the entire length of the helical hairpin of BCLxL and one face of the beta barrel of VDAC. Complex formation between BCL-xL and VDAC is also found by western blotting after cross-linking and size exclusion chromatography. These studies suggest that VDAC usually exists as a trimer (although oligomerization can be detergent dependent (Pappert and Schubert, 1983) and that within the trimer, BCL-xL binds to one or two VDACs, so that BCL-xL displaces one VDAC from a trimeric complex to form a new trimeric structure with BCL-xL as the third member. The data suggest that BCL-xL may change the structural properties of VDAC and enhance metabolite or cytochrome c conductance in different contexts. 6.2.3.3 BCL-xL Interaction with VDAC in Mitochondria Isolated from Ischemic Brain To prove that an interaction between BCL-xL and VDAC forms in vivo, a model system was studied in which application of recombinant BCL-xL protein to mitochondrial membranes produced channel activity that was attenuated by inhibiting VDAC (Jonas et al., 2004). BCL-xL is known to exist in two forms, a full-length antiapoptotic form and an N-truncated version (DN BCL-xL) that appears after death stimuli and acts as a cell killer protein (Clem et al., 1998). In mitochondria of synapses exposed to the death stimulus hypoxia or mitochondria isolated from ischemic brain, large channel activity is recorded that has biophysical features of the activity produced by application of DN BCL-xL to control mitochondria (Clem et al., 1998; Jonas et al., 2004; Bonanni et al., 2006). (Figs 6.3 and 6.5) Both the mitochondrial channel activity produced during hypoxia and the channel activated by the application of DN BCL-xL are strongly inhibited by millimolar concentrations of NADH. NADH binds with low

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affinity to VDAC, but in large concentrations can block its channel activity (Wunder and Colombini, 1991; Lee et al., 1994). NADH is ineffective on DN BCL-xL-induced channel activity when VDAC is absent (Basanez et al., 2001, 2002; Jonas et al., 2004). Furthermore, yeast mitochondria lacking VDAC (Lohret and Kinnally, 1995) do not respond to the application of recombinant DN BCL-xL with any change in channel activity, and NADH fails to inhibit the endogenous channel activity of the mitochondria lacking VDAC. These studies suggest that the large channel activity of mitochondria recorded after a death stimulus (hypoxia) is produced by a channel made up of a protein complex of VDAC and BCL-xL. Whether BCL-xL and VDAC actually interact biochemically or merely biophysically in vivo is still in question. 6.2.3.4 VDAC2 Inhibits Apoptosis In mammals, three different VDAC genes encoding distinct isoforms have been reported (Shoshan-Barmatz et al., 2006). More complex multicellular organisms have all three isoforms, suggesting that the different isoforms have specialized functions. Yeasts have two isoforms. Yeast VDAC1 protein forms pores when reconstituted into artificial lipid membranes, but yeast VDAC2 does not. VDAC1 and 3 are the predominant pore-forming isomers of VDAC in mammalian cells, but a role for VDAC2 has recently been suggested (Cheng et al., 2003). Cells deficient in VDAC2, but not VDAC1, are more susceptible to Bak oligomerization and apoptotic cell death, and these events are prevented by overexpression of VDAC2. In a model created to explain these findings, the authors suggest that the proapoptotic molecules tBID, BIM, or BAD may displace VDAC2 from Bak, enabling homooligo merization of Bak and release of cytochrome c through the mitochondrial outer membrane. 6.2.3.5 VDAC BAD Regulation of Metabolism and Cell Death The potential interaction of VDAC with hexokinase, as already described, places VDAC in a position to interact directly or indirectly with BAD, a BH3-only protein (Fig. 6.7). BAD is necessary in pancreas for the formation of a mitochondrial outer membrane complex of glucokinase (pancreatic hexokinase), protein kinase A, protein phosphatase 1, and Wiskott-Aldrich protein family member WAVE-1, a known actin binding protein. The BAD knockout animals or those in which BAD has been constitutively dephosphorylated display profound deficits in glucose homeostasis and glucose tolerance (Danial et al., 2003; Danial et al., 2008). In normal cultured cells deprived of glucose and in brain exposed to transient ischemia, BAD becomes dephosphorylated, translocates to mitochondrial membranes, and contributes to cell death, possibly by contributing to the formation of a cytochrome c-releasing pore (Fig. 6.7). Such a pore could be formed by the activation of Bax after sequestration of BCL-xL by BAD, or possibly by the activation of proapoptotic BCL-xL, after cleavage by caspases or calpains to form DN BCL-xL (Fig. 6.7) (Miyawaki et al., 2008). 6.2.4

VDAC Regulation of Learning and Memory

Studies of VDAC1 and 3 knockout mice suggest that VDAC plays an important role in the normal process of learning and memory (Weeber et al., 2002). Although there

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FIGURE 6.7 Physiological and pathological roles of BAD. In a normal pancreatic insulinsecreting cell, phosphorylated BAD nucleates a complex of proteins at the mitochondrial membrane that regulates normal glucose homeostasis. In BAD knockout animals (BAD
/
), the complex is disrupted and glucose metabolism is disturbed perhaps by a decrease in efficiency of ATP production in response to glucose stimulation. During cell death, BAD becomes dephosphorylated and participates in cytochrome c release from mitochondria, possibly by binding to BCL-xL and contributing to the formation of a channel produced by proapoptotic DN BCL-xL.

appears to be no obvious neuronal structural defect in the VDAC knockout mice, they are deficient in contextual fear conditioning and in the ability to learn to find a hidden platform in the Morris Water Maze test, suggesting that the mice lack normal capabilities for establishing memory and learning. In addition, hippocampal slice recordings from VDAC knockout mice demonstrate a decrease in certain forms of short-term facilitation. The cellular model proposed to underlie hippocampal memory formation and learning is long-term potentiation (LTP) of synaptic transmission. LTP is also significantly impaired in the knockout mice. Interestingly, low-dose CSA, which inhibits mitochondrial inner membrane depolarization, attenuates LTP in normal mice, suggesting that inner membrane ion channel activity is also involved in the onset of learning.

6.3 MITOCHONDRIAL INNER MEMBRANE CHANNELS Many studies of inner membrane physiology over the last half century were concerned with the management of calcium fluxes, calcium buffering by the matrix, and catastrophic calcium-induced depolarization of mitochondria (Nicholls and

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Akerman, 1982; Gunter and Pfeiffer, 1990; Bernardi, 1999). The next section will attempt to outline the inner membrane ion channel conductances as they relate to the mechanisms of calcium movement between the mitochondrion and the cytosol and the ability of the mitochondrion to produce energy in the form of ATP. Ion channels that regulate ion and metabolite fluxes across the inner membrane may form complexes with channels in the outer membrane such that the complex of channels may regulate the flow of ions and metabolites directly across the two membranes between the matrix and the cytosol. Regulation of complex formation may thereby alter metabolism and neuronal function. 6.3.1

Energy Dependence of Mitochondrial Calcium Accumulation

Calcium normally cycles constantly between the mitochondrial matrix and the cytosol. Calcium enters the matrix over its electrochemical gradient via the uniporter, a calcium-selective channel of the inner membrane (Litsky and Pfeiffer, 1997; Kirichok et al., 2004), and can be exchanged with sodium (sodium in, calcium out, Naþ/Ca2þ exchanger) linked to a sodium/hydrogen (Naþ/Hþ, hydrogen in, sodium out) exchanger and Hþ efflux via respiratory complexes (Fig. 6.8). These exchange pathways are used when cytosolic calcium levels are relatively low and no net accumulation of calcium occurs in the mitochondrial matrix (Fig. 6.8). Nevertheless, when calcium rises more precipitously in the cytosol, mitochondria can accumulate calcium and therefore may serve as major buffers for cytoplasmic calcium (Nicholls and Akerman, 1982). In this setting, Hþ ions are pumped out in 1. Calcium ci uniporter

Ca2+

3. Na +/H+ antiporter

5. Permeability transition pore

Ca2+ Na+

Na+

H+

2. Na +/Ca2+ exchanger

H+

H+

4. Electron transport complexes

H+

Ca2+ ADP

7. H +/Pi- cotransporter

ATP Pi

6. ATP synthase

CaPi

FIGURE 6.8 Management of calcium homeostasis in cells by mitochondrial calcium buffering. When cytosolic calcium levels are low, calcium cycles into and out of the matrix using channels and transporters (1–4). When cytosolic calcium levels rise, mitochondria can buffer calcium by the net exchange of calcium for Hþ ions, compromising the synthesis of ATP (6). The buffering of calcium is aided by the formation of a calcium phosphate complex in the matrix (7). Calcium can be rereleased through the transporters (2–4) or through a calciumactivated channel that may be related to the mPTP (5).

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exchange for calcium, so that net accumulation of calcium will occur together with net efflux of Hþ ions (Nicholls and Akerman, 1982). Therefore, the net accumulation of calcium leads to matrix alkalinization, and increases the difference in pH across the inner membrane. Eventually, no more Hþ can be lost from the matrix without compromising the maximum value of the membrane potential. When this happens, the membrane begins to depolarize toward 0 mV. More extensive calcium uptake can occur in the presence of anions (such as Pi, acetate, HCO3
) that permeate upon cotransport with Hþ ions (Fig. 6.7). The cotransport of Hþ and anions is stimulated by the previous alkalinization of the matrix. The resultant reestablishment of the normal concentration of Hþ ions in the matrix then allows the respiratory chain to reestablish the membrane potential by pumping out Hþ ions again, so that there may be only a transient membrane depolarization during calcium uptake. If phosphate is the permeating anion, then a calcium phosphate precipitate forms in the matrix, so that free calcium remains low. Under these conditions, there is theoretically possible a massive accumulation of precipitated calcium but very little accumulation of free matrix calcium. In the presence of phosphate, unlike with other anions such as acetate that do not form a precipitate, there is no change in osmolarity of the matrix. One Hþ accumulates in the cytosol for each calcium taken up, as the precipitate forms and Hþ is freed up from hydrogen phosphate. The increase in free Hþ in the matrix stimulates the respiratory complexes to pump Hþ back out by accelerating the respiratory chain to maintain the membrane potential, just as if ATP were being made. Hþ ions can also be pumped out by reversal of the synthase, requiring ATP hydrolysis. All of the above described processes take energy, as Hþ is pumped out but calcium flows in at the expense of ATP production. If no phosphate is available, then this reaction is prevented and further alkalinization of the matrix occurs with gradual depletion of the Hþ pool, finally resulting in depolarization of the membrane potential, preventing further calcium accumulation. 6.3.2

Voltage-Dependent Inner Membrane Channels: The Calcium Uniporter

Calcium enters the matrix via the uniporter, which appears to be a calcium selective, voltage-dependent channel (Kirichok et al., 2004). By recording in the whole mitoplast configuration, the problem of dissipation of the membrane potential by calcium influx can be prevented by voltage-clamp. In this mode, a calcium-selective current is increased by varying cytoplasmic calcium concentrations. The membrane is highly conductive to calcium under these circumstances, suggesting that the permeability pathway constitutes a channel rather than a transporter. At micromolar concentrations of calcium, the current density is comparable to that of voltage-gated plasma membrane calcium channels that are exposed to millimolar calcium. This is because of the large electrochemical potential gradient or driving force across the mitochondrial membrane and because the channel is usually in the open state. The calcium current is inwardly rectifying, so that it is larger at negative potentials, as one would expect for calcium uptake in energized mitochondria. The current is inactivated by time to a plateau level still carrying significant current. The total current is inhibited effectively by ruthenium

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red, a known inhibitor of calcium uniporter activity, but in single-channel recordings of inside-out patches, ruthenium is ineffective at positive potentials. In the absence of calcium, single-channel recordings demonstrate that the channel conducts sodium with a constant current but conducts calcium with a flickering current at positive potentials. The flickering in a calcium-containing medium suggests that the channel is partially blocked while calcium is passing through the channel. Although the molecular identity of the calcium uniporter is still not known, one candidate for calcium uptake in heart is the ryanodine receptor, which is activated by calcium and inhibited by Mg2þ and ruthenium red (Beutner et al., 2001). The ryanodine receptor may play an important role in the regulation of calcium uptake during cardiac ischemia. 6.3.3

Other Inner Membrane Conductances: Mitoplast Recording Technique

Patch clamping of isolated mitochondrial inner membranes in the “mitoplast” configuration has yielded data on several different conductances, but the most well studied is the large-conductance voltage-dependent channel of the inner membrane, variously named in the literature, but most likely serving as the underlying conductance of the rapid calcium efflux pathway related to the mitochondrial permeability transition (Crompton, 1999; Halestrap et al., 2000). The channel or pore that may underlie this conductance was termed mitochondrial permeability transition pore (mPTP). To the mPTPs have also been ascribed various functions in addition to calcium efflux, most notably a role in the formation of the inner membrane component of a two-membrane-spanning mitochondrial death channel complex that has been particularly implicated in ischemic cell death in heart and brain (Fig. 6.3) (Baines et al., 2005; Halestrap, 2005; Nakagawa et al., 2005). The first “mitoplast” recordings were obtained in the 1980s from isolated inner membranes from cuprizone-fed animals. Cuprizone feeding yields giant mitoplasts (5 mM or more in diameter) compared to those derived from control animals (1–3 mM in diameter), creating an organelle closer to the size of a small cell. This type of mitoplast is easier to patch clamp. The recordings (Sorgato et al., 1987, 1989) reveal a slightly anion-selective channel of 108 pS that is voltage dependent in that there is much more current, both whole “cell” or single channel, at positive potentials. 6.3.4 Channel Activity Correlated with Permeability Transition: The Mitochondrial Permeability Conductance Pore (mPTP) The permeability transition is characterized by a sudden loss of mitochondrial membrane potential induced by an increase in permeability of the inner membrane to a large number of unrelated solutes (Gunter and Pfeiffer, 1990). Physiologically relevant molecules and those for which there is no known influx pathway enter by diffusion down their concentration gradients, because after permeability transition, all energy-dependent processes of the inner membrane are halted by the dissipation of the potential gradient. The transition is regulated by calcium on the matrix side and therefore it serves as a calcium efflux pathway from the matrix to the cytosol, but whether this channel is used by mitochondria during physiological activity is still

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controversial. Swelling of the matrix also occurs after permeability transition because of the rapid influx of ions and small molecules followed by water. Swelling can be measured optically by the decrease in light scattering by the dissolved solutes. Therefore, permeability transition as measured with optical techniques can be correlated with channel activity as measured by patch-clamp recording of mitoplasts. The “MCC” has been characterized as a multiconductance, voltage-dependent channel recorded in mitoplast (inner membrane) preparations (Kinnally and Tedeschi, 1994) Most frequently, the activity was more prominent at negative potentials, with closings to lower conductances at positive potentials. The activity was found to be inhibited by amiodarone or cyclosporine A. These inhibitors caused the channel to step through various lower conductance levels and remain in the closed or small conductance open state for many minutes. The channel has biophysical characteristics in common with the permeability transition pore (mPTP) (Gunter and Pfeiffer, 1990) and is also known as mitochondrial megachannel (MMC) (Szabo and Zoratti, 1991, 1992; Bernardi et al., 1992). It is activated by calcium, inhibited by Mg2þ, and is voltage dependent (Kinnally et al., 1991; Bernardi, 1992; Bernardi et al., 1992; Scorrano et al., 1997). The similarities between the conductances recorded by different groups also includes inhibition by ADP and by alterations in pH. The MCC has the same response to various agents that inhibit the MMC, including propranolol, amiodarone, dibucaine, and benzodiazepines (Kinnally et al., 1992; Zorov et al., 1992). In another study performed at that time, a 1.3 nS conductance channel was recorded that also demonstrated subconductance states (Petronilli et al., 1989). The channel was voltage dependent. When the matrix was made positive, the channel was more likely to enter subconductance states. Unlike VDAC, at all negative potentials, the channel remained at a high conductance. It was hypothesized that these channels could participate in the transport of proteins across the inner membrane, but subsequent studies identified other more specific protein-conducting channels, therefore its complete function is still not fully understood. Further studies of the 1.3 nS channel revealed that it was activated by calcium on the matrix side and inhibited by competition of calcium with Mg2þ and Ba2þ, Sr2þ, and Mn2þ (Szabo et al., 1992). The calciumactivated 1.3 nS conductance was also inhibited by CSA, a known inhibitor of the prolyl isomerase cyclophilin, on the matrix side (Szabo and Zoratti, 1991). Cyclophilin had previously been known to inhibit permeability transition measured optically. The finding that channel activity was inhibited by the binding of CSA to cyclophilin established that cyclophilin was in a complex with the channel pore protein, whose molecular nature was and is still not completely known. Studies of cyclophilin D knockout mice by two independent groups (Baines et al., 2005; Nakagawa et al., 2005) subsequently established that cyclophilin D, and by inference the mPTP, was involved in ischemic necrosis, because mitochondria isolated from the knockout mice were resistant to calcium-induced and CSA-inhibited permeability transition. Correlated with the resistance to permeability transition was the resistance to necrotic death induced by ROS and calcium overload and to cardiac ischemia/reperfusion injury (Nakagawa et al., 2005). Tissues from animals overexpressing cyclophilin D had abnormal swollen mitochondria and a propensity toward spontaneous cell death (Baines et al., 2005). Despite these findings, developmental apoptotic death appeared

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normal. The mice underwent normal development, raising suspicions that this type of death did not require cyclophilin-regulated inner membrane channel activity. The prevalent hypothesis is that the mPTP may contain both inner and outer membrane components, such as the ANT that normally transports ATP out of the inner membrane, VDAC, and cyclophilin D (Fig. 6.3). The role of the ANT remains unproven, because in hepatocyte mitochondria isolated from knockout animals lacking both liver-expressed isoforms of ANT, calcium-induced permeability transition still occurs and is nonresponsive to inhibitors of ANT such as bongkrekic acid (Kokoszka et al., 2004). Interestingly, when isolated brain mitochondria are tested for their responses to calcium and CSA, full substrates allow full permeability transition in response to introduction of high external calcium but restricted substrates (such as succinate only) produce a partial depolarization that is sensitive to CSA but not sensitive to another mPTP inhibitor, bongkrekic acid, suggesting that ANT is not involved in this type of transition. Maximum respiratory stimulation is restricted by the low-level use of the electron transport chain in the setting of the restricted substrates, which enter the chain distal to Complex I, so that maximum Hþ pumping cannot be attained (Brustovetsky and Dubinsky, 2000). A regulation of substrate use by mitochondria may protect mitochondria from maximum respiratory stimulation and reactive oxygen species (ROS) production at times of calcium overload. Nevertheless, ATP production will still be limited by the partial depolarization and could be insufficient in times of stress (Brustovetsky and Dubinsky, 2000). 6.3.5 Physiological Function of the Two-Membrane-Spanning Channel Complex: Regulation of Contact Point Formation It is tempting to speculate that VDAC forms a molecular complex with an inner membrane channel, so that metabolites and ions may enter and exit the matrix directly from the cytosol (Fig. 6.3). Coimmunoprecipitation studies support this hypothesis suggesting that VDAC, ANT, and cyclophilin D are biochemically linked (Crompton et al., 1998). VDAC clusters at contact sites where inner membrane MCC-like activity is present by patch recording (Brdiczka et al., 1986; Sandri et al., 1988; Moran et al., 1990). There are estimated to be about 37 contact sites per square micron of mitochondrial membrane, making it likely that there are 7 contact sites in a recording of a membrane patch of 0.5 . Because of the importance of communication between the matrix and the cytosol, it is possible that a functional mitochondrial channel could require the presence of both inner and outer membrane components at contact sites in series (Halestrap, 2005). This would provide for streamlined flow of ions and metabolites from the matrix bypassing the intermembrane space and exiting directly into the cytosol. If this were to be the only type of conduit, however, there would be no way for intermembrane space components to be taken up or released through a channel. For example, cytochrome c, which resides in the intermembrane space, could not be released to the cytosol in the absence of outer membrane rupture (Brustovetsky et al., 2002; Green and Kroemer, 2004) Opposing the idea that channels of the two

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membranes exist in a fixed complex is the finding that recordings of MCC in yeast mitoplasts (with intact contact points) derived from cells lacking VDAC have channel activity that is similar to that of MCC recorded in mitoplasts from wild-type cells (Lohret and Kinnally, 1995), and that normal developmental apoptosis (with cytochrome c release) occurs in cyclophilin D knockout animals despite the lack of normal inner membrane channel regulation (Baines et al., 2005; Nakagawa et al., 2005). Nevertheless, contact point formation may be transient. For example, metabolic functions of contact sites have been proposed. A contact between the membranes may occur depending on the need to release ATP, to permit the entry of ADP into the matrix, or to support the entry of calcium. Contact sites may form when oxidative phosphorylation is taking place (Knoll and Brdiczka, 1983). In keeping with this idea, ADP, atractyloside, and succinate, a substrate for mitochondrial respiration, appear to induce contacts, whereas glycerol and uncouplers such as DNP or the electron transport inhibitor antimycin A decrease their formation (Hackenbrock, 1972; Bucheler et al., 1991). Contact sites isolated by differential centrifugation are distinct in densities, containing only outer or only inner membranes and may contain all the channel and regulatory proteins in the complex spanning the two membranes (Ohlendieck et al., 1986). Bonanni et al. 2006 have observed a decrease in electron density of contact sites after ischemia, suggesting that channel regulatory proteins may disperse from the complex, perhaps as a result of damage to the electron transport chain during the ischemic event. One possible explanation for the multiconductance state of many mitochondrial channels could be the ability to increase conductance depending on an increasing interaction between the two membranes (Kinnally and Tedeschi, 1994). Finally, it is possible that VDAC may transiently interact with members of the BCL2 family and hexokinase, either to regulate metabolism or during cell death (Figs 6.3 and 6.6), and this interaction may enhance or decrease the efficiency of mitochondrial metabolism, suggesting that complex formation may be regulated by events occurring outside of the outer membrane.

6.4 TRANSLOCATOR OF THE INNER MEMBRANE (TIM) AND TRANSLOCATOR OF THE OUTER MEMBRANE (TOM) The protein translocators of the outer and inner mitochondrial membranes are waterfilled channels that provide pathways for mitochondrial proteins translated in the cytosol in the form of preproteins encoded by nuclear genes. Eight different tranlocator proteins of the outer membrane have been identified that participate in preprotein translocation, recognition of preproteins, and insertion of resident outer membrane proteins (Kunkele et al., 1998) and these eight proteins function as receptors and/or pore-forming subunits. Tom40 and Tom22 form the major constituents of the Tom complex (Kunkele et al., 1998). Tom40 is an integral membrane protein with mainly beta sheet structure that forms a cation-selective high conductance in artificial lipid bilayer recordings. It binds to mitochondrial targeting sequences added to the cis side of the membrane. The whole Tom complex forms a single-ring structure of 75 Awith a pore size of 16–20 A. Tom40 forms channels that are mostly open below 100 mV,

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with occasional, flickering closures and prominent subconductance states at negative potentials. The main conductance of Tom40 in 250 mM KCl is 360 pS, with a subconductance state of 150 pS and a cation over anion selectivity of 8:1. The current–voltage relationship shows voltage dependence with the greatest probability of opening near 0 and symmetrically increased tendency toward closure as voltage is increased on both sides of 0 mV. The channel is transiently blocked by mitochondrial preproteins and mitochondrial signal peptides, but not by related peptides lacking the mitochondrial signal (Hill et al., 1998). Upon direct comparison of the single-channel properties of Tom and Tim, both channels were found to demonstrate high conductance, voltage dependence, and slight cation selectivity. The channels were similarly affected by signal sequence peptides (Muro et al., 2003).

6.5 MITOCHONDRIAL CHANNEL CONDUCTANCE: EFFECTS ON SYNAPTIC STRENGTH Mitochondria are the predominant organelle within many presynaptic terminals. During frequent synaptic events, they affect intracellular calcium homeostasis and provide the energy needed for synaptic vesicle recycling and for the continued operation of membrane ion pumps. Recent discoveries have altered our ideas about the role of mitochondria in the synapse. Mitochondrial localization, morphology, and docking at synaptic sites may indeed alter the kinetics of transmitter release and calcium homeostasis in the presynaptic terminal. In addition, mitochondrial ion channel activity of BCL-xL alters synaptic transmitter release and the recycling of pools of synaptic vesicles. BCL-xL, therefore, not only affects the life and death of the cell soma, but its actions in the synapse may underlie the regulation of basic synaptic processes that subtend learning, memory, and synaptic development. Mitochondrial ion channels participate in the management of cytosolic calcium levels and in the release of ATP and are therefore potentially extremely important for the regulation of synaptic transmission (Blaustein et al., 1978). Different types of neuronal synapses contain different numbers of mitochondria with slightly different properties, depending on whether the main function of the mitochondria is to provide energy or buffer calcium. In most synapses, oxidative metabolism by mitochondria is crucial to successful neurotransmission (Nguyen et al., 1997). Moreover, mitochondrial bioenergetics are altered acutely in synapses that have undergone preconditioning, providing for enhanced oxidative competence (Nguyen et al., 1997), suggesting that an interaction may exist between neuronal plasticity and mitochondrial plasticity (Nguyen and Atwood, 1994). 6.5.1

Mitochondria Alter Calcium Homeostasis During Synaptic Events

Synaptic transmission depends on mitochondria not only for energy production, but also for maintaining calcium homeostasis within the presynaptic terminal (Kaftan et al., 2000; Nicholls and Budd, 2000; Atwood and Karunanithi, 2002; Jonas, 2006). During synaptic events, calcium influx through voltage-gated channels and the

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release of calcium from intracellular stores including mitochondria produce elevations of cytosolic calcium that enhance synaptic vesicle fusion (Stevens, 2003). In the crayfish neuromuscular junction, modulation of fast synaptic transmission is dependent on persistently elevated calcium levels inside the presynaptic terminal produced by mitochondrial calcium rerelease (Tang and Zucker, 1997) Different synapses may have different degrees of potentiation or depression of release of neurotransmitter, depending on their ability to buffer and make calcium available at certain times (Wang and Kaczmarek, 1998; Billups and Forsythe, 2002; Stevens, 2003). The proximity of sites of calcium influx to sites of vesicle fusion also contributes to enhancing the probability of fusion events (Sakaba and Neher, 2001; Schneggenburger et al., 2002). In some synapses, reaccumulation of vesicles to different pools that have been depleted may be differentially dependent on ATP (Jonas, 2006). Release of calcium from mitochondria participates in shaping the time course and amplitude of neurotransmitter release from presynaptic nerve endings after the invasion of the endings by action potentials. In the example of the crayfish neuromuscular junction, eliminating the ability of mitochondria to sequester calcium not only leads to a higher rise in cytosolic calcium inside the presynaptic terminal during a tetanus, but also to prevention of the normal potentiation of neurotransmitter release after the tetanus (Tang and Zucker, 1997). 6.5.2 Mitochondrial Presence at Presynaptic Sites Regulates Intense Synaptic Activity Mitochondrial ATP release also appears to play a role in the management of vesicle pool size both at rest and during synaptic stimulation. In Drosophila melanogaster, mitochondrial targeting is necessary for normal synaptic transmission at the neuromuscular junction (Stowers et al., 2002; Guo et al., 2005; Verstreken et al., 2005). Animals lacking mitochondrial targeting proteins such as Milton (Stowers et al., 2002), GTPase dMiro (Guo et al., 2005), or syntabulin (Cai et al., 2005) cannot target mitochondria normally to the presynaptic terminal and have abnormal synaptic transmission. Animals lacking normal mitochondrial division, which regulates the targeting of mitochondria to developing synapses, also have abnormal synapses (Li et al., 2004; Verstreken et al., 2005). 6.5.3 Mitochondrial ATP Production Regulates Normal Functioning of Synaptic Vesicle Pools Distinct pools of vesicles have different probabilities of release (Rizzoli and Betz, 2005). The readily releasable pool is defined as the vesicles that are immediately available for release, or “docked” at the active zone. In hippocampal synapses, for example, there appear to be approximately 5–10 vesicles that are docked at each active zone, but a single brief stimulus (such as an action potential) may release only one vesicle. The recycling pool is defined as the pool of vesicles that continue to be released and reaccumulate during moderate or physiological stimulation. This pool contains

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FIGURE 6.9 Regulation of mitochondrial ATP release enhances synaptic transmission, in part by regulating the refilling of depleted neurotransmitter-containing vesicle pools.

5–20% of all vesicles, but these estimates vary in different synapses. The reserve pool is defined as those vesicles that only release upon extremely frequent stimulation. The reserve pool of vesicles makes up about 80–90% of the vesicles in most terminals. The temperature-sensitive Drosophila shibire mutant (Kuromi and Kidokoro, 2002) demonstrates that the reserve pool of vesicles is normally mobilized only after the recycling pool is depleted. This mutant exhibits defective endocytosis at high temperatures, leading to an inability of vesicles to reaccumulate after exocytosis. In conditions of mild or moderate stimulation, which would not usually mobilize the reserve pool in controls, the reserve pool is mobilized at high temperatures in the mutant because it cannot refill its recycling pool. The experiments suggest that the reserve pool will be used only after the recycling pool has been depleted. The recycling pool, therefore, may contain vesicles that are privileged for release, either by their interaction with specific cytoskeletal elements, or their location, or both (Rizzoli and Betz, 2005). Surprisingly, however, the recycling pool is not located adjacent to the active zone. Rather, the vesicles of the recycling pool are distributed widely throughout the vesicle cluster (Rizzoli and Betz, 2004). ATP is required for certain steps in synaptic vesicle mobilization, release, and recycling. Specific ATP-dependent steps in synaptic transmission include refilling single vesicles with neurotransmitter (Takamori et al., 2000), membrane fission during endocytosis (Heidelberger, 2001), and coated pit formation (Smythe et al., 1989; Faundez and Kelly, 2000). Recent evidence suggests that ATP is required for normal functioning of vesicle pools (Kuromi and Kidokoro, 2002; Verstreken et al., 2005). Studies of the Drp1 mutation in Drosophila suggest that mobilizing the reserve pool requires ATP. These synapses do not contain mitochondria. An ATP-sensitive motor, the mysosin light chain kinase, which moves vesicles from pool to pool in an energydependent manner, is affected by the lack of locally released ATP brought on by the absence of mitochondria at synaptic endings. It is clear that mitochondria need to be targeted to the synapse for synaptic transmission to function normally during intense stimulation. How does the release of ATP from mitochondria increase at the time it is needed during intense stimulation?

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As seen in electron micrographs, the brainstem auditory synapse of the medial nucleus of the trapezoid body (MNTB) (the Calyx of Held), which is specialized to release neurotransmitter at extremely high frequency and fidelity, contains a mitochondrial adherens complex. The complex is a collection of filaments that tether mitochondria very closely to the synapse in a regulated fashion, orienting the matrix cristae perpendicular to the active zone (Rowland et al., 2000). It is likely that the organization of mitochondria within this specialized synapse enables the mitochondria to carry out precisely timed ATP release and calcium buffering. In hippocampal neurons, which have considerably different synaptic organization than that observed in the Calyx of Held, it appears that mitochondria are mostly untethered and that they can be either mobile or stationary. When hippocampal neurons are stimulated by local application of growth factors to points on the axon, mitochondria move preferentially to the stimulated site, presumably mimicking the in vivo situation where mitochondria might be targeted rapidly during growth or plasticity (Chada and Hollenbeck, 2003, 2004; Malaiyandi et al., 2005). 6.5.4

Mitochondrial Ion Channel Regulation During Synaptic Transmission

Specific targeting of mitochondria is thus required for normal synaptic transmission at high frequencies. The regulated targeting of mitochondria to sites of high energy demand suggests that the mechanisms of ATP production and release by mitochondria could very well be regulated during frequent synaptic events. Mitochondria are suggested to release ATP via VDAC, as described above. It is predicted that during synaptic events (such as synaptic plasticity), regulation of the opening of VDAC in the outer mitochondrial membrane could occur. Another prediction is that there is likely to be a second messenger that signals the opening of VDAC during synaptic events. The first evidence that mitochondrial ion channel activity could be regulated during synaptic events came from studies of mitochondrial membrane conductance during synaptic transmission in an intact presynaptic terminal of the squid stellate ganglion. Through the use of a double-barreled patch pipette (Jonas et al., 1997), recordings were made both at rest and during and after intense synaptic stimulation (Jonas et al., 1999). In control recordings within the resting squid presynaptic terminal, the conductance of mitochondrial membranes is low. In contrast, during frequent electrical stimulation of the squid presynaptic nerve, there occurs up to a 60-fold increase in activity and conductance of mitochondrial membrane patches within the presynaptic terminal (Jonas et al., 1999), a change that lasts for approximately 1 min after the stimulus. The delay and persistence of the mitochondrial membrane activity after stimulation implies that the mitochondrial outer membrane channel activity is not simultaneous with the opening of plasma membrane channels and suggests that the increase depends on an intracellular second messenger. Such a messenger could be calcium, which remains elevated in the squid terminal for approximately 1 min after stimulation, just as in the crayfish neuromuscular junction (Jonas, 2006). In keeping with these reports, in a calcium-deficient bathing medium, there is no change in mitochondrial conductance in response to stimulation of the presynaptic terminal, demonstrating that the evoked

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mitochondrial membrane channel activity is dependent on calcium influx into the terminal (Jonas et al., 1999). In addition, the uncoupler FCCP (carbonyl cyanide p-trifluoromethoxyphenylhydrazone), which depolarizes mitochondria, prevents the channel activity. The acute changes in mitochondrial membrane activity are correlated with synaptic plasticity, because FCCP application eliminates the short-term potentiation of the synapse that follows nerve stimulation. The findings imply that a calcium-dependent channel such as the mPTP might be involved in the change in conductance. The calcium-dependent site of that protein complex is in the matrix, however, and the recordings were performed on the outer mitochondrial membrane, suggesting that the activity could represent opening of an inner membrane channel whose activity might be linked to the opening of VDAC or another conductance in the outer membrane (Halestrap, 2005). A channel spanning two membranes could, therefore, permit the efflux of calcium (as well as ATP and other ions and metabolites) from the matrix into the cytosol during synaptic potentiation. In addition to mPTP components such as VDAC and ANT, BCL-xL and other BCL2 family members might also contribute to the regulation of channel activity by the protein complex during synaptic transmission. As we have seen, BCL-xL resides in the outer membranes of mitochondria (Kaufmann et al., 2003) and the recombinant protein induces mitochondrial ion channel activity (Jonas et al., 2003). BCL-xL could influence the release of calcium, ATP, or other metabolites into the cytosol during synaptic responses. In support of this hypothesis, injection of recombinant BCL-xL protein into the presynaptic terminal enhances the rate of rise of postsynaptic responses, resulting in an earlier latency for evoked action potentials in the postsynaptic cell as compared to the latency recorded in control presynaptic termini (Jonas et al., 2003). Interestingly, the injected BCL-xL protein produces potentiation of synaptic transmitter release in both healthy synapses and in those in which transmission had run down. Under these conditions, injection of BCL-xL protein into the terminal enhances the amplitude of the postsynaptic potential, restoring suprathreshold responses, in effect, giving the synapse a “new life.” If BCL-xL regulates the flux of metabolites across the outer mitochondrial membrane (Vander Heiden et al., 2001; Gottlieb et al., 2002), then this property predicts that the regulation of levels of ATP may enhance neurotransmission in the physiological setting. The evidence to support this hypothesis comes from the studies of the effect of ATP injection into the synapse on the degree of synaptic responses (Jonas et al., 2003; Verstreken et al., 2005). Direct microinjection of ATP into the synapse produces a similar degree and time course of enhancement of synaptic transmission as the effects of BCL-xL injection (Jonas et al., 2003) and, in fact, occludes the effects of injection of BCL-xL, suggesting that the two agents acted via the same mechanism. The findings imply that the conductance change could involve a calcium-dependent increase in the activity of ATP release regulated by BCL-xL. Although the injection of recombinant BCL-xL protein enhances transmission for a prolonged period, endogenous BCL-xL might participate in short-term responses of the mitochondrial membrane after high intensity stimulation. Recent findings suggest that this is indeed the case. The activities of BCL-xL can be disrupted by the application of ABT-737, a mimetic of the BH3-only protein BAD, that binds to BCL-xL with high

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affinity within a pocket of the three-dimensional structure that usually binds proapoptotic BH3-only proteins (Oltersdorf et al., 2005). The structure of ABT-737, a thioethylamino-2-4-dimethylphenyl analogue, was designed to bind to the threedimensional structure of BCL-xL/BCL-2. It displaces a GFP-tagged BH3-only protein from BCL-xL at mitochondrial surfaces in intact tumor cells. In cancer cell lines, ABT737 alone effectively induces cell death possibly via its ability to displace the prebound proapoptotic proteins Bax and Bak from BCL-xL (Oltersdorf et al., 2005). When applied to mitochondria within the squid presynaptic terminal just before synaptic transmission, ABT-737 inhibits the channel activity of mitochondrial membranes induced by synaptic stimulation, suggesting that BCL-xL is necessary for this activity. Recovery of vesicle pools after synaptic depression is also regulated by BCL-xL. Stimulation of the synapse at 2 Hz produces synaptic depression as the readily releasable pool is rapidly depleted (Swandulla et al., 1991). After the depletion, the more reluctantly releasable pools are accessed. During the recovery phase following the administration of a tetanus given against the background of continuous 2 Hz stimulation, the vesicles do not repopulate the readily releasable pool (because of the continued 2 H3 stimulation) but do repopulate the reluctant pools. The time course of the recovery of the reluctant pools is rapid (Sakaba and Neher, 2001) and is not affected by the previous injection of BCL-xL. If recovery from tetanic stimulation is measured during infrequent basal stimulation, full recovery of all pools occurs after a tetanus, as evidenced by the ability of the synapse to release as fully after the tetanus as it does during the control period at the beginning of the experiment. Nevertheless, the time course of the recovery of synaptic responses following the tetanus is slower than it is at 2 Hz, suggesting that when all the pools are repopulated, the most readily releasable––the first pool to be released at the onset of stimulation––repopulates quite slowly (Sakaba and Neher, 2001; Sakaba et al., 2005). The amount of recovery of this pool is significantly enhanced by BCL-xL injection when compared to recovery measured in controls. Thus, a slow component of the time course of recovery of the total vesicle pool is sensitive to the actions of BCLxL, and the pool that is affected may be the most readily releasable pool. BCL-xL appears to enhance the ability of this subset of neurotransmitter-containing vesicles to become available for release. If endogenous BCL-xL is necessary for recovery of synaptic vesicle pools, then ABT-737 might affect the rate of recovery from tetanic stimulation. In control squid synapses, recovery of neurotransmitter release after a tetanus generally occurs in less than 2 min. In contrast, in synapses exposed to ABT-737 just before tetanic stimulation, the rate at which the synapse recovers from high frequency firing is decreased (Hickman et al., 2008). As we have seen, the enhancement of synaptic responses by FL BCL-xL could be related to a difference in function of its ion channel compared to that of proapoptotic molecules such as Bax or DNBCL-xL. A key characteristic of the ion channel activity of BCL-xL is that it can induce ATP exchange across mitochondrial membranes (Vander Heiden et al., 2000, 2001). In particular, it performs this function in mitochondria from cells that have been exposed to apoptotic stimuli such as growth factor deprivation. In this pathological setting, BCL-xL may protect cells from death by

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maintaining ATP levels in the cell and by keeping VDAC in its open configuration. The delicate balance between the pro- and antiapoptotic BCL-2 family proteins may thereby regulate mitochondrial metabolism at times of stress and may control the onset of the eventual release of proapoptotic mitochondrial factors into the cytosol (Plas and Thompson, 2002). These factors and the compromise of mitochondrial function caused by the release of these factors may together cause the decline in synaptic responses. A surprising dichotomy of the effects of antiapoptotic molecules is that they may enhance the release of ATP from mitochondria (Gottlieb et al., 2002), but prevent the release of cytochrome c (Antonsson et al., 1997; Kluck et al., 1997). Binding of BCLxL and BCL-2 to proapoptotic molecules may inhibit channel activity of the proapoptotic molecules. Therefore, the channel activity of FL BCL-xL as well as its ability to alter the channel activities of prodeath molecules may comprise the antiapoptotic functions of FL BCL-xL. In summary, although large conductance mitochondrial membrane activity causes cell death and synaptic dysfunction, smaller conductance activity may be a prerequisite for a normal and healthy neuronal life.

ACKNOWLEDGMENTS This work was supported by NIH RO1 NS045876 and an American Heart Established Investigator Award (EAJ).

REFERENCES Adams JM, Cory S, 2007. The Bcl-2 apoptotic switch in cancer development and therapy. Oncogene 26: 1324–1337. Antonsson B, Conti F, Ciavatta A, Montessuit S, Lewis S, Martinou I, Bernasconi L, Bernard A, Mermod JJ, Mazzei G, Maundrell K, Gambale F, Sadoul R, Martinou JC, 1997. Inhibition of Bax channel-forming activity by Bcl-2. Science 277: 370–372. Antonsson B, Montessuit S, Lauper S, Eskes R, Martinou JC, 2000. Bax oligomerization is required for channel-forming activity in liposomes and to trigger cytochrome c release from mitochondria. Biochem J 345(Pt 2): 271–278. Atwood HL, Karunanithi S, 2002. Diversification of synaptic strength: presynaptic elements. Nat Rev Neurosci 3: 497–516. Baines CP, Kaiser RA, Purcell NH, Blair NS, Osinska H, Hambleton MA, Brunskill EW, Sayen MR, Gottlieb RA, Dorn GW, Robbins J, Molkentin JD, 2005. Loss of cyclophilin D reveals a critical role for mitochondrial permeability transition in cell death. Nature 434: 658–662. Baines CP, Kaiser RA, Sheiko T, Craigen WJ, Molkentin JD, 2007. Voltage-dependent anion channels are dispensable for mitochondrial-dependent cell death. Nat Cell Biol 9: 550–555. Banasiak KJ, Xia Y, Haddad GG, 2000. Mechanisms underlying hypoxia-induced neuronal apoptosis. Prog Neurobiol 62: 215–249. Basanez G, Zhang J, Chau BN, Maksaev GI, Frolov VA, Brandt TA, Burch J, Hardwick JM, Zimmerberg J, 2001. Pro-apoptotic cleavage products of Bcl-xL form cytochrome cconducting pores in pure lipid membranes. J Biol Chem 276: 31083–31091.

184

UNIQUE MITOCHONDRIAL ION CHANNELS

Basanez G, Sharpe JC, Galanis J, Brandt TB, Hardwick JM, Zimmerberg J, 2002. Bax-type apoptotic proteins porate pure lipid bilayers through a mechanism sensitive to intrinsic monolayer curvature. J Biol Chem 277: 49360–49365. Bernardi P, 1992. Modulation of the mitochondrial cyclosporin A-sensitive permeability transition pore by the proton electrochemical gradient evidence that the pore can be opened by membrane depolarization. J Biol Chem 267: 8834–8839. Bernardi P, 1999. Mitochondrial transport of cations: channels, exchangers, and permeability transition. Physiol Rev 79: 1127–1155. Bernardi P, Vassanelli S, Veronese P, Colonna R, Szabo I, Zoratti M, 1992b. Modulation of the mitochondrial permeability transition pore effect of protons and divalent cations. J Biol Chem 267: 2934–2939. Beutner G, Sharma VK, Giovannucci DR, Yule DI, Sheu SS, 2001. Identification of a ryanodine receptor in rat heart mitochondria. J Biol Chem 276: 21482–21488. Billups B, Forsythe ID, 2002. Presynaptic mitochondrial calcium sequestration influences transmission at mammalian central synapses. J Neurosci 22: 5840–5847. Blachly-Dyson E, Peng SZ, Colombini M, Forte M, 1989. Probing the structure of the mitochondrial channel, VDAC, by site-directed mutagenesis: a progress report. J Bioenerg Biomembr 21: 471–483. Blachly-Dyson E, Peng S, Colombini M, Forte M, 1990. Selectivity changes in site-directed mutants of the VDAC ion channel: structural implications. Science 247: 1233–1236. Blaustein MP, Ratzlaff RW, Kendrick NK, 1978. The regulation of intracellular calcium in presynaptic nerve terminals. Ann N Y Acad Sci 307: 195–212. Bonanni L, Chachar M, Jover-Mengual T, Li H, Jones A, Yokota H, Ofengeim D, Flannery RJ, Miyawaki T, Cho CH, Polster BM, Pypaert M, Hardwick JM, Sensi SL, Zukin RS, Jonas EA, 2006. Zinc-dependent multi-conductance channel activity in mitochondria isolated from ischemic brain. J Neurosci 26: 6851–6862. Brdiczka D, Knoll G, Riesinger I, Weiler U, Klug G, Benz R, Krause J, 1986. Microcompartmentation at the mitochondrial surface: its function in metabolic regulation. Adv Exp Med Biol 194: 55–69. Brdiczka D, Kaldis P, Wallimann T, 1994. In vitro complex formation between the octamer of mitochondrial creatine kinase and porin. J Biol Chem 269: 27640–27644. Brustovetsky N, Dubinsky JM, 2000. Dual responses of CNS mitochondria to elevated calcium. J Neurosci 20: 103–113. Brustovetsky N, Brustovetsky T, Jemmerson R, Dubinsky JM, 2002. Calcium-induced cytochrome c release from CNS mitochondria is associated with the permeability transition and rupture of the outer membrane. J Neurochem 80: 207–218. Bucheler K, Adams V, Brdiczka D, 1991. Localization of the ATP/ADP translocator in the inner membrane and regulation of contact sites between mitochondrial envelope membranes by ADP. A study on freeze-fractured isolated liver mitochondria. Biochim Biophys Acta 1056: 233–242. Cai Q, Gerwin C, Sheng ZH, 2005. Syntabulin-mediated anterograde transport of mitochondria along neuronal processes. J Cell Biol 170: 959–969. Chada SR, Hollenbeck PJ, 2003. Mitochondrial movement and positioning in axons: the role of growth factor signaling. J Exp Biol 206: 1985–1992. Chada SR, Hollenbeck PJ, 2004. Nerve growth factor signaling regulates motility and docking of axonal mitochondria. Curr Biol 14: 1272–1276.

REFERENCES

185

Cheng EH, Kirsch DG, Clem RJ, Ravi R, Kastan MB, Bedi A, Ueno K, Hardwick JM, 1997. Conversion of Bcl-2 to a Bax-like death effector by caspases. Science 278: 1966–1968. Cheng EH, Sheiko TV, Fisher JK, Craigen WJ, Korsmeyer SJ, 2003. VDAC2 inhibits BAK activation and mitochondrial apoptosis. Science 301: 513–517. Clem RJ, Cheng EH, Karp CL, Kirsch DG, Ueno K, Takahashi A, Kastan MB, Griffin DE, Earnshaw WC, Veliuona MA, Hardwick JM, 1998. Modulation of cell death by Bcl-XL through caspase interaction. Proc Natl Acad Sci USA 95: 554–559. Colombini M, 1989. Voltage gating in the mitochondrial channel VDAC. J Membr Biol 111: 103–111. Colombini M, Yeung CL, Tung J, Konig T, 1987. The mitochondrial outer membrane channel, VDAC, is regulated by a synthetic polyanion. Biochim Biophys Acta 905: 279–286. Colombini M, Blachly-Dyson E, Forte M, 1996. VDAC, a channel in the outer mitochondrial membrane. Ion Channels 4: 169–202. Crompton M, 1999. The mitochondrial permeability transition pore and its role in cell death. Biochem J 341: 233–249. Crompton M, Virji S, Ward JM, 1998. Cyclophilin-D binds strongly to complexes of the voltage-dependent anion channel and the adenine nucleotide translocase to form the permeability transition pore. Eur J Biochem 258: 729–735. Danial NN, Gramm CF, Scorrano L, Zhang CY, Krauss S, Ranger AM, Datta SR, Greenberg ME, Licklider LJ, Lowell BB, Gygi SP, Korsmeyer SJ, 2003. BAD and glucokinase reside in a mitochondrial complex that integrates glycolysis and apoptosis. Nature 424: 952– 956. Danial NN, Walensky LD, Zhang CY, Choi CS, Fisher JK, Molina AJ, Datta SR, Pitter KL, Bird GH, Wikstrom JD, Deeney JT, Robertson K, Morash J, Kulkarni A, Neschen S, Kim S, Greenberg ME, Corkey BE, Shirihai OS, Shulman GI, Lowell BB, Korsmeyer SJ, 2008. Dual role of proapoptotic BAD in insulin secretion and beta cell survival. Nat Med 14: 144– 153. Dejean LM, Martinez-Caballero S, Guo L, Hughes C, Teijido O, Ducret T, Ichas F, Korsmeyer SJ, Antonsson B, Jonas EA, Kinnally KW, 2005. Oligomeric Bax is a component of the putative cytochrome c release channel MAC, mitochondrial apoptosis-induced channel. Mol Biol Cell 16: 2424–2432. Denis-Pouxviel C, Riesinger I, Buhler C, Brdiczka D, Murat JC, 1987. Regulation of mitochondrial hexokinase in cultured HT 29 human cancer cells. An ultrastructural and biochemical study. Biochim Biophys Acta 902: 335–348. Doring C, Colombini M, 1985. Voltage dependence and ion selectivity of the mitochondrial channel, VDAC, are modified by succinic anhydride. J Membr Biol 83: 81–86. Fannjiang Y, Kim CH, Huganir RL, Zou S, Lindsten T, Thompson CB, Mito T, Traystman RJ, Larsen T, Griffin DE, Mandir AS, Dawson TM, Dike S, Sappington AL, Kerr DA, Jonas EA, Kaczmarek LK, Hardwick JM, 2003. BAK alters neuronal excitability and can switch from anti- to pro-death function during postnatal development. Dev Cell 4: 575–585. Faundez VV, Kelly RB, 2000. The AP-3 complex required for endosomal synaptic vesicle biogenesis is associated with a casein kinase I alpha-like isoform. Mol Biol Cell 11: 2591– 2604. Galonek HL, Hardwick JM, 2006. Upgrading the BCL-2 network. Nat Cell Biol 8: 1317–1319. (See comment)

186

UNIQUE MITOCHONDRIAL ION CHANNELS

Golshani-Hebroni SG, Bessman SP, 1997. Hexokinase binding to mitochondria: a basis for proliferative energy metabolism. J Bioenerg Biomembr 29: 331–338. Gottlieb E, Armour SM, Thompson CB, 2002. Mitochondrial respiratory control is lost during growth factor deprivation. Proc Natl Acad Sci USA 99: 12801–12806. Green DR, Kroemer G, 2004. The pathophysiology of mitochondrial cell death. Science 305: 626–629. Gunter TE, Pfeiffer DR, 1990. Mechanisms by which mitochondria transport calcium. Am J Physiol 258: C755–C786. Guo X, Macleod GT, Wellington A, Hu F, Panchumarthi S, Schoenfield M, Marin L, Charlton MP, Atwood HL, Zinsmaier KE, 2005. The GTPase dMiro is required for axonal transport of mitochondria to Drosophila synapses. Neuron 47: 379–393. Hackenbrock CR, 1972a. Energy-linked ultrastructural transformations in isolated liver mitochondria and mitoplasts. Preservation of configurations by freeze-cleaving compared to chemical fixation. J Cell Biol 53: 450–465. Hackenbrock CR, 1972. States of activity and structure in mitochondrial membranes. Ann N Y Acad Sci 195: 492–505. Halestrap A, 2005. Biochemistry: a pore way to die. Nature 434: 578–579. (See comment) Halestrap AP, Doran E, Gillespie JP, O’Toole A, 2000. Mitochondria and cell death. Biochem Soc Trans 28: 170–177. Hashimoto M, Wilson JE, 2000. Membrane potential-dependent conformational changes in mitochondrially bound hexokinase of brain. Arch Biochem Biophys 384: 163–173. Heidelberger R, 2001. ATP is required at an early step in compensatory endocytosis in synaptic terminals. J Neurosci 21: 6467–6474. Hickman JA, Hardwick JM, Kaczmarek LK, Jonas EA, 2008. BcL-xL inhibitor ABT-737 reveals a dual role for BcL-xL in synaptic transmission. J Neurophysiol 99: 1515–1522. Hill K, Model K, Ryan MT, Dietmeier K, Martin F, Wagner R, Pfanner N, 1998. Tom40 forms the hydrophilic channel of the mitochondrial import pore for preproteins. Nature 395: 516– 521. Hodge T, Colombini M, 1997. Regulation of metabolite flux through voltage-gating of VDAC channels. J Membr Biol 157: 271–279. Holden MJ, Colombini M, 1988. The mitochondrial outer membrane channel, VDAC, is modulated by a soluble protein. FEBS Lett 241: 105–109. Holden MJ, Colombini M, 1993. The outer mitochondrial membrane channel, VDAC, is modulated by a protein localized in the intermembrane space. Biochim Biophys Acta 1144: 396–402. Hsu YT, Youle RJ, 1998. Bax in murine thymus is a soluble monomeric protein that displays differential detergent-induced conformations. J Biol Chem 273: 10777–10783. Jonas E, 2006. BCL-xL regulates synaptic plasticity. Mol Interventions 6: 208–222. Jonas EA, Knox RJ, Kaczmarek LK, 1997. Giga-ohm seals on intracellular membranes: a technique for studying intracellular ion channels in intact cells. Neuron 19: 7–13. Jonas EA, Buchanan J, Kaczmarek LK, 1999. Prolonged activation of mitochondrial conductances during synaptic transmission. Science 286: 1347–1350. Jonas EA, Hoit D, Hickman JA, Brandt TA, Polster BM, Fannjiang Y, McCarthy E, Montanez MK, Hardwick JM, Kaczmarek LK, 2003. Modulation of synaptic transmission by the BCL2 family protein BCL-xL. J Neurosci 23: 8423–8431.

REFERENCES

187

Jonas EA, Hickman JA, Chachar M, Polster BM, Brandt TA, Fannjiang Y, Ivanovska I, Basanez G, Kinnally KW, Zimmerberg J, Hardwick JM, Kaczmarek LK, 2004. Proapoptotic Ntruncated BCL-xL protein activates endogenous mitochondrial channels in living synaptic terminals. Proc Natl Acad Sci USA 101: 13590–13595. Jonas EA, Hardwick JM, Kaczmarek LK, 2005. Actions of BAX on mitochondrial channel activity and on synaptic transmission. Antioxid Redox Signal 7: 1092–1100. Kaftan EJ, Xu T, Abercrombie RF, Hille B, 2000. Mitochondria shape hormonally induced cytoplasmic calcium oscillations and modulate exocytosis. J Biol Chem 275: 25465– 25470. Kaufmann T, Schlipf S, Sanz J, Neubert K, Stein R, Borner C, 2003. Characterization of the signal that directs Bcl-x(L), but not Bcl-2, to the mitochondrial outer membrane. J Cell Biol 160: 53–64. Kim H, Rafiuddin-Shah M, Tu HC, Jeffers JR, Zambetti GP, Hsieh JJ, Cheng EH, 2006. Hierarchical regulation of mitochondrion-dependent apoptosis by BCL-2 subfamilies Nat Cell Biol 8: 1348–1358. Kinnally KW, Tedeschi H, 1994. Mitochondrial channels: an integrated view. In: Forte M, Colombini M (Eds), Molecular Biology of Mitochondrial Transport Systems. SpringerVerlag, Berlin, pp. 169–198. Kinnally KW, Zorov D, Antonenko Y, Perini S, 1991. Calcium modulation of mitochondrial inner membrane channel activity. Biochem Biophys Res Commun 176: 1183–1188. Kinnally KW, Antonenko YN, Zorov DB, 1992. Modulation of inner mitochondrial membrane channel activity. J Bioenerg Biomembr 24: 99–110. Kirichok Y, Krapivinsky G, Clapham DE, 2004. The mitochondrial calcium uniporter is a highly selective ion channel. Nature 427: 360–364. Kluck RM, Bossy-Wetzel E, Green DR, Newmeyer DD, 1997. The release of cytochrome c from mitochondria: a primary site for Bcl-2 regulation of apoptosis. Science 275: 1132– 1136. Knoll G, Brdiczka D, 1983. Changes in freeze-fractured mitochondrial membranes correlated to their energetic state. Dynamic interactions of the boundary membranes. Biochim Biophys Acta 733: 102–110. Kokoszka JE, Waymire KG, Levy SE, Sligh JE, Cai J, Jones DP, MacGregor GR, Wallace DC, 2004. The ADP/ATP translocator is not essential for the mitochondrial permeability transition pore. Nature 427: 461–465. Kroemer G, Reed JC, 2000. Mitochondrial control of cell death. Nat Med 6: 513–519. Kunkele KP, Heins S, Dembowski M, Nargang FE, Benz R, Thieffry M, Walz J, Lill R, Nussberger S, Neupert W, 1998. The preprotein translocation channel of the outer membrane of mitochondria. Cell 93: 1009–1019. Kuromi H, Kidokoro Y, 2002. Selective replenishment of two vesicle pools depends on the source of Ca2þ at the Drosophila synapse. Neuron 35: 333–343. Lee AC, Zizi M, Colombini M, 1994. Beta-NADH decreases the permeability of the mitochondrial outer membrane to ADP by a factor of 6. J Biol Chem 269: 30974–30980. Lemasters JJ, Holmuhamedov E, 2006. Voltage-dependent anion channel (VDAC) as mitochondrial governator—thinking outside the box. Biochim Biophys Acta 1762: 181–190. Lewis J, Oyler GA, Ueno K, Fannjiang YR, Chau BN, Vornov J, Korsmeyer SJ, Zou S, Hardwick JM, 1999. Inhibition of virus-induced neuronal apoptosis by Bax. Nat Med 5: 832–835.

188

UNIQUE MITOCHONDRIAL ION CHANNELS

Li Z, Okamoto K, Hayashi Y, Sheng M, 2004. The importance of dendritic mitochondria in the morphogenesis and plasticity of spines and synapses. Cell 119: 873–887. Litsky ML, Pfeiffer DR, 1997. Regulation of the mitochondrial Ca2þ uniporter by external adenine nucleotides: the uniporter behaves like a gated channel which is regulated by nucleotides and divalent cations. Biochemistry (Mosc) 36: 7071–7080. Lohret TA, Kinnally KW, 1995. Multiple conductance channel activity of wild-type and voltage-dependent anion-selective channel (VDAC)-less yeast mitochondria. Biophys J 68: 2299–2309. Malaiyandi LM, Honick AS, Rintoul GL, Wang QJ, Reynolds IJ, 2005. Zn2þ inhibits mitochondrial movement in neurons by phosphatidylinositol 3-kinase activation. J Neurosci 25: 9507–9514. Malia TJ, Wagner G, 2007. NMR structural investigation of the mitochondrial outer membrane proteinVDAC and itsinteraction withantiapoptotic Bcl-xL.Biochemistry(Mosc)46:514–525. Mangan PS, Colombini M, 1987. Ultrasteep voltage dependence in a membrane channel. Proc Natl Acad Sci USA 84: 4896–4900. Martinez-Caballero S, Dejean LM, Jonas EA, Kinnally KW, 2005. The role of the mitochondrial apoptosis induced channel MAC in cytochrome c release. J Bioenerg Biomembr 37: 155–164. Middleton G, Davies AM, 2001. Populations of NGF-dependent neurones differ in their requirement for BAX to undergo apoptosis in the absence of NGF/TrkA signalling in vivo. Development 128: 4715–4728. Minn AJ, Velez P, Schendel SL, Liang H, Muchmore SW, Fesik SW, Fill M, Thompson CB, 1997. Bcl-x(L) forms an ion channel in synthetic lipid membranes. Nature 385: 353–357. Miyawaki T, Mashiko T, Ofengeim D, Flannery RJ, Noh KM, Fujisawa S, Bonanni L, Bennett MV, Zukin RS, Jonas EA, 2008. Ischemic preconditioning blocks BAD translocation, BcL-xL cleavage, and large channel activity in mitochondria of postischemic hippocampal neurons. Proc Natl Acad Sci USA 105: 4892–4897. Moran O, Sandri G, Panfili E, Stuhmer W, Sorgato MC, 1990. Electrophysiological characterization of contact sites in brain mitochondria. J Biol Chem 265: 908–913. Muchmore SW, Sattler M, Liang H, Meadows RP, Harlan JE, Yoon HS, Nettesheim D, Chang BS, Thompson CB, Wong SL, Ng SL, Fesik SW, 1996. X-ray and NMR structure of human Bcl-xL, an inhibitor of programmed cell death. Nature 381: 335–341. Muro C, Grigoriev SM, Pietkiewicz D, Kinnally KW, Campo ML, 2003. Comparison of the TIM and TOM channel activities of the mitochondrial protein import complexes. Biophys J 84: 2981–2989. Nakagawa T, Shimizu S, Watanabe T, Yamaguchi O, Otsu K, Yamagata H, Inohara H, Kubo T, Tsujimoto Y, 2005. Cyclophilin D-dependent mitochondrial permeability transition regulates some necrotic but not apoptotic cell death. Nature 434: 652–658. Nguyen PV, Atwood HL, 1994. Altered impulse activity modifies synaptic physiology and mitochondria in crayfish phasic motor neurons. J Neurophysiol 72: 2944–2955. Nguyen PV, Marin L, Atwood HL, 1997. Synaptic physiology and mitochondrial function in crayfish tonic and phasic motor neurons. J Neurophysiol 78: 281–294. Nicholls D, Akerman K, 1982. Mitochondrial calcium transport. Biochim Biophys Acta 683: 57–88. Nicholls DG, Budd SL, 2000. Mitochondria and neuronal survival. Physiol Rev 80: 315–360.

REFERENCES

189

Ohlendieck K, Riesinger I, Adams V, Krause J, Brdiczka D, 1986. Enrichment and biochemical characterization of boundary membrane contact sites from rat-liver mitochondria. Biochim Biophys Acta 860: 672–689. Oltersdorf T, Elmore SW, Shoemaker AR, Armstrong RC, Augeri DJ, Belli BA, Bruncko M, Deckwerth TL, Dinges J, Hajduk PJ, Joseph MK, Kitada S, Korsmeyer SJ, Kunzer AR, Letai A, Li C, Mitten MJ, Nettesheim DG, Ng S, Nimmer PM, O’Connor JM, Oleksijew A, Petros AM, Reed JC, Shen W, Tahir SK, Thompson CB, Tomaselli KJ, Wang B, Wendt MD, Zhang H, Fesik SW, Rosenberg SH, 2005. An inhibitor of Bcl-2 family proteins induces regression of solid tumours. Nature 435: 677–681. Pappert G, Schubert D, 1983. The state of association of band 3 protein of the human erythrocyte membrane in solutions of nonionic detergents. Biochim Biophys Acta 730: 32–40. Pavlov EV, Priault M, Pietkiewicz D, Cheng EH, Antonsson B, Manon S, Korsmeyer SJ, Mannella CA, Kinnally KW, 2001. A novel, high conductance channel of mitochondria linked to apoptosis in mammalian cells and Bax expression in yeast. J Cell Biol 155: 725–731. Peng S, Blachly-Dyson E, Forte M, Colombini M, 1992. Large scale rearrangement of protein domains is associated with voltage gating of the VDAC channel. Biophys J 62: 123–131. Penso J, Beitner R, 1998. Clotrimazole and bifonazole detach hexokinase from mitochondria of melanoma cells. Eur J Pharmacol 342: 113–117. Petronilli V, Szabo I, Zoratti M, 1989. The inner mitochondrial membrane contains ionconducting channels similar to those found in bacteria. FEBS Lett 259: 137–143. Plas DR, Thompson CB, 2002. Cell metabolism in the regulation of programmed cell death. Trends Endocrin Metab 13: 75–78. Polster BM, Kinnally KW, Fiskum G, 2001. BH3 death domain peptide induces cell-type selective mitochondrial outer membrane permeability. J Biol Chem 276: 37887– 37894. Rizzoli SO, Betz WJ, 2004. The structural organization of the readily releasable pool of synaptic vesicles. Science 303: 2037–2039. Rizzoli SO, Betz WJ, 2005. Synaptic vesicle pools. Nat Rev Neurosci 6: 57–69. Rostovtseva TK, Antonsson B, Suzuki M, Youle RJ, Colombini M, Bezrukov SM, 2004. Bid, but not Bax, regulates VDAC channels. J Biol Chem 279: 13575–13583. Rostovtseva TK, Tan W, Colombini M, 2005. On the role of VDAC in apoptosis: fact and fiction. J Bioenerg Biomembr 37: 129–142. Roucou X, Montessuit S, Antonsson B, Martinou JC, 2002. Bax oligomerization in mitochondrial membranes requires tBid (caspase-8-cleaved Bid) and a mitochondrial protein. Biochem J 368: 915–921. Rowland KC, Irby NK, Spirou GA, 2000. Specialized synapse-associated structures within the calyx of Held. J Neurosci 20: 9135–9144. Sakaba T, Neher E, 2001. Calmodulin mediates rapid recruitment of fast-releasing synaptic vesicles at a calyx-type synapse. Neuron 32: 1119–1131. Sakaba T, Stein A, Jahn R, Neher E, 2005. Distinct kinetic changes in neurotransmitter release after SNARE protein cleavage. Science 309: 491–494. Sandri G, Siagri M, Panfili E, 1988. Influence of Ca2þ on the isolation from rat brain mitochondria of a fraction enriched of boundary membrane contact sites. Cell Calcium 9: 159–165. Schendel SL, Montal M, Reed JC, 1998. Bcl-2 family proteins as ion channels. Cell Death Differ 5: 372–380.

190

UNIQUE MITOCHONDRIAL ION CHANNELS

Schlesinger PH, Gross A, Yin XM, Yamamoto K, Saito M, Waksman G, Korsmeyer SJ, 1997. Comparison of the ion channel characteristics of proapoptotic BAX and antiapoptotic BCL2. Proc Natl Acad Sci USA 94: 11357–11362. Schneggenburger R, Sakaba T, Neher E, 2002. Vesicle pools and short-term synaptic depression: lessons from a large synapse. Trends Neurosci 25: 206–212. Scorrano L, Petronilli V, Bernardi P, 1997. On the voltage dependence of the mitochondrial permeability transition pore. A critical appraisal. J Biol Chem 272: 12295–12299. Shimizu S, Narita M, Tsujimoto Y, 1999. Bcl-2 family proteins regulate the release of apoptogenic cytochrome c by the mitochondrial channel VDAC. Nature 399: 483–487. (See comment; erratum appears in Nature 2000, 407(6805):767) Shimizu S, Shinohara Y, Tsujimoto Y, 2000. Bax and Bcl-xL independently regulate apoptotic changes of yeast mitochondria that require VDAC but not adenine nucleotide translocator. Oncogene 19: 4309–4318. Shoshan-Barmatz V, Israelson A, Brdiczka D, Sheu SS, 2006. The voltage-dependent anion channel (VDAC): function in intracellular signalling, cell life and cell death. Curr Pharm Des 12: 2249–2270. Smythe E, Pypaert M, Lucocq J, Warren G, 1989. Formation of coated vesicles from coated pits in broken A431 cells. J Cell Biol 108: 843–853. Sorgato MC, Keller BU, Stuhmer W, 1987. Patch-clamping of the inner mitochondrial membrane reveals a voltage-dependent ion channel. Nature 330: 498–500. Sorgato MC, Moran O, De Pinto V, Keller BU, Stuehmer W, 1989. Further investigation on the high-conductance ion channel of the inner membrane of mitochondria. J Bioenerg Biomembr 21: 485–496. Stevens CF, 2003. Neurotransmitter release at central synapses. Neuron 40: 381–388. Stowers RS, Megeath LJ, Gorska-Andrzejak J, Meinertzhagen IA, Schwarz TL, 2002. Axonal transport of mitochondria to synapses depends on milton, a novel Drosophila protein. Neuron 36: 1063–1077. Sugioka R, Shimizu S, Funatsu T, Tamagawa H, Sawa Y, Kawakami T, Tsujimoto Y, 2003. BH4domain peptide from Bcl-xL exerts anti-apoptotic activity in vivo. Oncogene 22: 8432–8440. Swandulla D, Hans M, Zipser K, Augustine GJ, 1991. Role of residual calcium in synaptic depression and posttetanic potentiation: fast and slow calcium signaling in nerve terminals. Neuron 7: 915–926. Szabo I, Zoratti M, 1991. The giant channel of the inner mitochondrial membrane is inhibited by cyclosporin A. J Biol Chem 266: 3376–3379. Szabo I, Zoratti M, 1992. The mitochondrial megachannel is the permeability transition pore. J Bioenerg Biomembr 24: 111–117. Szabo I, Bernardi P, Zoratti M, 1992. Modulation of the mitochondrial megachannel by divalent cations and protons. J Biol Chem 267: 2940–2946. Takamori S, Rhee JS, Rosenmund C, Jahn R, 2000. Identification of a vesicular glutamate transporter that defines a glutamatergic phenotype in neurons. Nature 407: 189–194. Tang Y, Zucker RS, 1997. Mitochondrial involvement in post-tetanic potentiation of synaptic transmission. Neuron 18: 483–491. Tsujimoto Y, Shimizu S, 2000. VDAC regulation by the Bcl-2 family of proteins. Cell Death Differ 7: 1174–1181. Tsujimoto Y, Cossman J, Jaffe E, Croce CM, 1985. Involvement of the Bcl-2 gene in human follicular lymphoma. Science 228: 1440–1443.

REFERENCES

191

Vander Heiden MG, Chandel NS, Li XX, Schumacker PT, Colombini M, Thompson CB, 2000. Outer mitochondrial membrane permeability can regulate coupled respiration and cell survival. Proc Natl Acad Sci USA 97: 4666–4671. Vander Heiden MG, Li XX, Gottleib E, Hill RB, Thompson CB, Colombini M, 2001. Bcl-xL promotes the open configuration of the voltage-dependent anion channel and metabolite passage through the outer mitochondrial membrane. J Biol Chem 276: 19414–19419. Verstreken P, Ly CV, Venken KJ, Koh TW, Zhou Y, Bellen HJ, 2005. Synaptic mitochondria are critical for mobilization of reserve pool vesicles at Drosophila neuromuscular junctions. Neuron 47: 365–378. Wang LY, Kaczmarek LK, 1998. High-frequency firing helps replenish the readily releasable pool of synaptic vesicles. Nature 394: 384–388. Weeber EJ, Levy M, Sampson MJ, Anflous K, Armstrong DL, Brown SE, Sweatt JD, Craigen WJ, 2002. The role of mitochondrial porins and the permeability transition pore in learning and synaptic plasticity. J Biol Chem 277: 18891–18897. Wicker U, Bucheler K, Gellerich FN, Wagner M, Kapischke M, Brdiczka D, 1993. Effect of macromolecules on the structure of the mitochondrial inter-membrane space and the regulation of hexokinase. Biochim Biophys Acta 1142: 228–239. Wolter KG, Hsu YT, Smith CL, Nechushtan A, Xi XG, Youle RJ, 1997. Movement of Bax from the cytosol to mitochondria during apoptosis. J Cell Biol 139: 1281–1292. Wood DE, Newcomb EW, 2000. Cleavage of Bax enhances its cell death function. Exp Cell Res 256: 375–382. Wunder UR, Colombini M, 1991. Patch clamping VDAC in liposomes containing whole mitochondrial membranes. J Membr Biol 123: 83–91. Youle RJ, Strasser A, 2008. The BCL-2 protein family: opposing activities that mediate cell death. Nat Rev Mol Cell Biol 9: 47–59. Yuan J, Yankner BA, 2000. Apoptosis in the nervous system. Nature 407: 802–809. Zalk R, Israelson A, Garty ES, Azoulay-Zohar H, Shoshan-Barmatz V, 2005. Oligomeric states of the voltage-dependent anion channel and cytochrome c release from mitochondria. Biochem J 386: 73–83. Zha J, Harada H, Yang E, Jockel J, Korsmeyer SJ, 1996. Serine phosphorylation of death agonist BAD in response to survival factor results in binding to 14-3-3 not BCL-X(L). Cell 87: 619–628. Zheng Y, Shi Y, Tian C, Jiang C, Jin H, Chen J, Almasan A, Tang H, Chen Q, 2004. Essential role of the voltage-dependent anion channel (VDAC) in mitochondrial permeability transition pore opening and cytochrome c release induced by arsenic trioxide. Oncogene 23: 1239–1247. Zimmerberg J, Parsegian VA, 1986. Polymer inaccessible volume changes during opening and closing of a voltage-dependent ionic channel. Nature 323: 36–39. Zizi M, Forte M, Blachly-Dyson E, Colombini M, 1994. NADH regulates the gating of VDAC, the mitochondrial outer membrane channel. J Biol Chem 269: 1614–1616. Zizi M, Byrd C, Boxus R, Colombini M, 1998. The voltage-gating process of the voltagedependent anion channel is sensitive to ion flow. Biophys J 75: 704–713. Zorov DB, Kinnally KW, Perini S, Tedeschi H, 1992. Multiple conductance levels in rat heart inner mitochondrial membranes studied by patch clamping. Biochim Biophys Acta 1105: 263–270. Zukin RS, Jover T, Yokota H, Calderone A, Simionescu M, Lau C-Y, 2004. Molecular and cellular mechanisms of ischemia-induced neuronal death. In: Mohr JP, Choi DW, Grotta JC, Weir B, Wolf PA (Eds), Stroke: Pathophysiology, Diagnosis, and Management. Churchill Livingstone, Philadelphia, pp. 829–854.

7 REGULATION OF NEURONAL EXCITABILITY BY THE SODIUM-ACTIVATED POTASSIUM CHANNELS SLICK (SLO2.1) AND SLACK (SLO2.2) VALENTIN K. GRIBKOFF1,2 AND LEONARD K. KACZMAREK2 1

Discovery Research, Knopp Neurosciences Inc., 2100 Wharton Street, Suite 615, Pittsburgh, PA 15203, USA 2 Department of Pharmacology, Yale University School of Medicine, New Haven, CT 06520, USA

7.1 INTRODUCTION Various chapters in this book describe neuronal voltage-gated ion channels that are regulated by intracellular messengers, including other ions. This chapter describes a subfamily of potassium (Kþ) channels, the Slo2 channels Slick and Slack, which encode the sodium (Naþ)-activated Kþ channels (KNa). As the name implies, these Kþ channels are regulated by the levels of intracellular Naþ ([Naþ]in) as well as by other ions and second messengers. Like Ca2þ, Naþ enters cells by multiple pathways, including specific voltage-gated ion channels. Unlike Ca2þ, which is an important second messenger in many cell types, Naþ currents have important roles principally in excitable cells, and therefore rapid changes or large, long-lasting changes in [Naþ]in are most prevalent in these cells. KNa channels are, therefore, not as widely distributed outside of the nervous system and muscle cells as Ca2þ-activated Kþ channels

Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

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(including Slo1, the BKCa channel; see Chapters 12 and 15), although with the cloning of KNa channels, the presence of KNa subunits can be more precisely determined and they are being localized to additional areas. In neurons, Naþ plays a pivotal role in the intercellular communication, and its intracellular concentration varies significantly as a function of cellular activity. Neuronal KNa channels have evolved to respond to activity-dependent increases in intracellular Naþ, and can act as a brake on the longer term effects of rapid or prolonged action potential discharge resulting from intracellular Naþ accumulation. 7.2 SODIUM-ACTIVATED POTASSIUM CURRENT (KNa) 7.2.1

KNa in Cardiac Myocytes

The first recordings made from KNa channels were not made in neurons, but rather in isolated guinea pig ventricular myocytes (Kameyama et al., 1984). The channels were of large conductance (>200 pS in 150 mM Kþ outside/49 mM Kþ inside, among the largest conductance cardiac ion channels), inwardly rectifying near the Kþ reversal potential but outwardly rectifying at more depolarized potentials, and activated by intracellular Naþ in excess of 20 mM with an EC50 in excess of 60 mM. The channels were not affected by varying ATP levels or pH within a physiological range, nor were they activated by intracellular Ca2þ. Some of these original findings, such as the pH sensitivity of the channels, have been challenged (Veldkamp et al., 1994), and such discrepancies may reflect the molecular makeup of different KNa channels. Importantly, the high threshold of activation of these channels by intracellular Naþ, in cells where the “normal” concentration is no greater than 10 mM, suggested that the current would primarily be activated under conditions in which Naþ extrusion from these myocytes was compromised. This could be due to failure of the Naþ-Kþ pump, a condition that could also result in deregulation of intracellular Ca2þ concentration when Naþ accumulated to the point of failure of the Naþ–Ca2þ exchange mechanism. A condition in which these criteria for Naþ-dependent opening of the channel would be met is cardiac hypoxia (Kameyama et al., 1984; Bertrand et al., 1989; Dryer, 1994; Veldkamp et al., 1994). In ventricular myocytes, this apparent low sensitivity of KNa to intracellular Naþ has been observed consistently, with activation occurring only at Naþ concentrations greater than 20–30 mM such as during block of the Naþ–Kþ-pump (Luk and Carmeliet, 1990; Rodrigo and Chapman, 1990; Niu and Meech, 2000). While some activation of KNa has been reported following physiological activation during which intracellular [Naþ]in is increased only by about 30% above the normal concentration of approximately 7 mM (Rodrigo, 1993), overall these data support the contention that the primary activation of this current in myocytes would occur during pathological conditions. In neurons, however, the sensitivity of KNa to intracellular Naþ may be higher and physiological increases in [Naþ]in are often much larger than in the myocytes. An early and consistent finding with native KNa channels from ventricular cells has been the existence of multiple, regularly spaced subconductance states (Sanguinetti, 1990; Wang et al., 1991; Mistry et al., 1996). Subconductance states, representing

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the fractional opening of a channel, can have a number of potential underlying mechanisms, although most evidence suggests that the structures of voltage-gated ion channels provide opportunities for multiple partial gating conformations (Fox, 1987; Dani and Fox, 1991; Bezanilla, 2005; Chapman and VanDongen, 2005). Subconductance states are observed in many ion channels, including ligandgated ion channels, particularly in the presence of compounds or experimental conditions that result in partial channel block or incomplete activation, or at membrane potentials where the probability of channel opening is small (Moss and Moczydlowski, 1996; Bezanilla, 2005; Chapman and VanDongen, 2005). Wang et al (1991) examined the conductance of KNa channels in ventricular myocytes and using variance-mean analysis (Patlak, 1993), found at least 10 major conductance sublevels. Interestingly, they found that these channels displayed inward rectification with outward current reduced by increasing intracellular Mg2þ, Naþ, or Ba2þ, but the appearance of subconductance states was apparently unaffected by the direction of the current (inward versus outward) or the presence of outward current inhibition by Mg2þ, Naþ, or Ba2þ. Subsequent analysis has shown that ventricular KNa channels may have 12 subconductance levels during longer, stable recordings, divided into transient and stable subconductances (Mistry et al., 1996). There were two stable levels, which were about 30 and 70% of the conductance of the fully open channel and which occurred spontaneously and for only relatively brief periods during prolonged recordings. The kinetics of the two observed stable subconductance levels, their open and closed times under identical conditions, were approximately equal to those of full conductance transitions when they were present (in other words, they look pretty much like smaller versions of the full open state when they occur, while the transient levels have very different kinetics with only brief open times). KNa channels in ventricular myocytes, therefore, activate in a highly complex manner, with at least 10 transient subconductance levels that likely contribute little in terms of physiological effect and at least 2 stable subconductance levels that occur spontaneously but that thus far largely defy experimental induction. These data strongly suggest that there are multiple stable channel configurations of native KNa channels. Additional characteristics of peripheral KNa channels have been described. These include pH sensitivity (block by Hþ, as mentioned above) (Veldkamp et al., 1994) and inhibition by Kþ (Niu and Meech, 2000). KNa currents and channels were recorded from neurons soon after their discovery in myocytes (Bertrand et al., 1989; Haimann and Bader, 1989; Martin and Dryer, 1989; Haimann et al., 1990; Kubota and Saito, 1991). In the following section we will briefly describe neuronal KNa channels and their functions, the identification of the genes encoding KNa channels, and the beginning of the pharmacological characterization of these channels. 7.2.2

KNa and the Regulation of Neuronal Excitability

Neurons successfully fulfill their numerous roles in sensation, information processing, secretion, and motor control in part because of their ability to change their response repertoire as a function of prior input and activity. Many neuronal constituents contribute to neuronal plasticity (used here in this broadest sense), including a number

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of ion channels. Both Ca2þ and Naþ are charge carriers and hence important mediators of neuronal function, each with their own set of ion channels that specifically control the flux of these cations across the plasma membrane (see Chapters 1 – 3). Ca2þ also serves as an important second messenger in many cells, and its intracellular concentration is tightly regulated. Since changes in the intracellular concentrations of both cations can have significant neuronal consequences, one or more corresponding Kþ channels have evolved to indirectly regulate their intracellular concentration and mitigate the effects that increases in their intracellular concentrations can have on neuronal excitability. There are two subfamilies of Kþ channels, the largeconductance Ca2þ-activated Kþ channels (BKCa, maxi-K, KCa1.1) encoded by the Slo1 gene (Butler et al., 1993; Dworetzky et al., 1996; Salkoff et al., 2006) and the small-and intermediate-conductance Ca2þ-activated Kþ channels KCa2.1–2.3 and KCa3.1 (Kohler et al., 1996; Ishii et al., 1997; Joiner et al., 1997; Bond et al., 1999) that respond to changes in [Ca2þ]in. Similarly, KNa channels have evolved in neurons to regulate excitability changes that result from intracellular Naþ accumulation (molecular identity to be discussed below). Importantly, unlike in cardiac myocytes, there is increasing evidence that KNa plays an important role in the normal regulation of neuronal activity, particularly in neurons where high-frequency action potential discharge activity is an important physiological feature. This has suggested that activation of KNa in neurons differs from that in myocytes in several possible ways, each of which have been or are currently being approached experimentally. For example, neuronal KNa channels may not require the high levels of [Naþ]in required by KNa in cardiac myocytes (where [Naþ]in only reaches activating levels during pathological conditions such as ischemia), or the levels of [Naþ]in in neurons may reach activating levels (>10 mM) during normal neuronal activity, at least in critical neuronal microenvironments. Both of these possibilities may be true at different times; KNa channels may be modulated by intracellular messengers, which may affect responsiveness to [Naþ]in, and quite high levels of [Naþ]in can be reached in select loci following prolonged neuronal activity, particularly after high-frequency discharge. In particular, the membrane hyperpolarizations following single and multiple action potentials contribute to the determination of action potential frequency and the adaptation of frequency (the reduction in firing observed during prolonged action potential trains) that are critical to the function of many neurons (Gean and ShinnickGallagher, 1989; Faber and Sah, 2005; Gu et al., 2007; Khawaja et al., 2007). These afterhyperpolarizations (AHPs) generally have multiple components. They vary significantly between different neurons in terms of underlying ionic mechanism, duration, and influence on cellular function, but Kþ currents initiated by action potentials or their sequelae (voltage changes or increased [Ca2þ]in or [Naþ]in) are wellcharacterized mediators of these events (Kim and McCormick, 1998; Wikstrom and El, 1998; Nedergaard, 2004; Pedarzani et al., 2005; Wallen et al., 2007). KNa has been shown to be an important contributor to AHPs, with increasing influence during prolonged and high-frequency neuronal activity. Finally, KNa may also serve a role during neuronal ischemia that is similar to that postulated for KNa in cardiac cells. Native KNa channels have been recorded from neurons in many areas in the central nervous system (CNS) and in the periphery. In the vertebrate CNS, KNa channels have

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been recorded from neocortical neurons, sensory neurons, hippocampal neurons, neurons within the auditory nuclei, spinal cord, and the olfactory system, among others (Bertrand et al., 1989; Haimann and Bader, 1989; Haimann et al., 1990, 1992; Egan et al., 1992a, 1992b; Dryer, 1994; Safronov and Vogel, 1996; Franceschetti et al., 2003; Liu and Stan Leung, 2004; Hess et al., 2007; Yang et al., 2007). In the olfactory bulb, KNa channels may be one of the most prominent and frequently recorded Kþ channels (Egan et al., 1992b). KNa channels represent one of the major mechanisms for responding to neuronal activity, particularly to prolonged neuronal activity at high frequencies, and the effect of activation of these channels is to produce a hyperpolarization and a decrease in membrane resistance. By the late 1980s/early 1990s, KNa channels exhibiting all of the hallmark characteristics of cardiac KNa channels, such as sensitivity to high [Naþ]in, inward rectification due to Naþ at highly positive membrane potentials, channel rundown in excised patches, and resistance to Liþ substitution for Naþ, had been recorded from central neurons. In several CNS areas, the role of KNa has been particularly well documented. As examples of this, we will briefly review some of the data concerning KNa functions in neurons of the neocortex and hippocampus and in neurons in the auditory system (specifically the medial nucleus of the trapezoid body (MNTB)). The purpose is not to exhaustively review this literature, but to introduce the function of KNa in neurons. A number of studies have examined the roles of these channels in regulating the excitability of neocortical and hippocampal neurons. In cat sensorimotor cortex, early studies demonstrated the existence and importance of KNa in regulating the responsiveness of large layer V neurons. When Ca2þ influx was blocked to remove the stimulus for Ca2þ-activated Kþ currents, a slow outward current resulted either from evoking prolonged discharges or by injecting subthreshold depolarizing current sufficient to activate persistent Naþ current (Schwindt et al., 1989). The outward current was blocked or greatly reduced in amplitude when persistent Naþ current was blocked using tetrodotoxin, and it produced spike frequency adaptation that could be reduced by certain G-protein-coupled receptors, such as norepinephrine (Schwindt et al., 1989). In visual cortical neurons, exposure to high-contrast visual stimuli results in neuronal adaptation. This is defined as a progressive loss of neuronal responsiveness to the continued presence of the stimulus, a phenomenon that is mediated at the cellular level (and revealed in intracellular current clamp recordings) by a long-lasting membrane hyperpolarization that is reflected in the degree of firing rate reduction in these cells (Sanchez-Vives et al., 2000a, 2000b). As mentioned above, similar spike frequency adaptation has been recorded in many types of neurons and it can have several underlying mechanisms. Sanchez-Vives and colleagues have shown, using ferret primary visual cortical slices in vitro, that prolonged depolarizing current injections or prolonged sinusoidal currents injected at a frequency of 2 Hz produced variable slow spike frequency adaptation over many seconds in these neurons. The discharge was followed by a prolonged AHP that lasted many seconds and was mediated by one or more Kþ currents. While the block of voltage-gated Ca2þ currents reduced the rate of spike frequency adaptation, the AHP was not affected. The reduction in [Naþ]out, the ultimate source of [Naþ]in during prolonged neuronal activity, however, further reduced the remaining spike frequency adaptation and also

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significantly reduced the AHP (Sanchez-Vives et al., 2000a). KNa currents therefore may be important, along with KCa currents, in tuning the responses of visual cortical neurons to persistent stimuli and play a critical role in such phenomena as contrast adaptation. A role for KNa can also be demonstrated for fast-spiking neurons of the visual cortex, cells that are capable of discharging at rates up to
1 kHz without adaptation and without significant changes in spike amplitude. In these neurons, the action potential duration is typically brief, the Naþ channels recover from inactivation rapidly, and the action potentials are short because these cells express voltagedependent Kþ currents (KV3 family) that activate rapidly and produce transient current. Together these factors allow the generation of high-frequency trains of action potentials. In these cells, brief periods of stimulation do not result in spike frequency adaptation, but a recent study demonstrated that longer trains of high-frequency action potentials did induce a slow AHP of many seconds duration, an effect that reduced the cells’ responses to afferents for much of this period (Descalzo et al., 2005). This slow AHP had all the characteristics of a KNa-mediated membrane hyperpolarization, although KNa channels were not definitively identified as underlying the AHP. The importance of KNa in neuronal regulation in cortical and hippocampal neurons can also be demonstrated in intrinsically bursting neurons. As opposed to neurons that discharge primarily or exclusively in response to supra-threshold extrinsic stimuli, the intrinsically bursting neurons in cortex and hippocampus have conductances that initiate and terminate bursts of action potentials in the absence of synaptic influences (Franceschetti et al., 1993; Istvan and Zarzecki, 1994; Metherate and Aramakis, 1999; Sipila et al., 2006). Identifying and understanding these properties has clear relevance to cortical function, as well as dysfunction, as exemplified by their involvement in epileptiform activity (Traub and Wong, 1982; Silva-Barrat et al., 1992; Sanabria et al., 2001). In sensorimotor cortical neurons in vitro, intrinsic bursts are followed by AHPs that are key determinants of burst length and rhythmic discharge. In these cells, partial substitution of Naþ with Liþ had little effect on action potentials (since voltagedependent Naþ channels will conduct Liþ current), but significantly increased burst duration and reduced AHP amplitude, an effect that was also observed on the Kþ conductance evoked following the Naþ entry through Naþ channels (Franceschetti et al., 2003). The effect of Liþ substitution was not observed when voltage-dependent Naþ channels were blocked with tetrodotoxin. Thus, KNa currents, which are reduced on substitution of Naþ by Liþ, appear to play a significant role in AHP generation and in the burst rhythmicity in neocortical neurons. The influence of KNa is not limited to reducing excitability during and following the prolonged trains of action potentials or bursts. In hippocampal pyramidal neurons, the depolarizing afterpotentials (DAPs) are important regulators of neuronal excitability and can be produced by single-antidromic or current-evoked action potentials. The replacement of [Naþ]out by Liþ enhanced DAP amplitudes, produced spike bursts where single-action potentials had been evoked, and caused a positive shift in the reversal potential of the DAP (Liu and Stan Leung, 2004). These data suggest that even after a single-action potential, KNa can influence the subsequent excitability of central neurons. In the auditory system, the binaural pathway in the brainstem permits the localization of the source of sounds by comparing the time and intensity differences

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of sound stimuli received at the two cochlea. To accomplish this with high fidelity, the neurons in various nuclei, such as the neurons of the anterior ventral cochlear nucleus and their targets among the principal neurons of the MNTB, are capable of maintaining action potential discharge rates faithfully locked to the phase of incoming synaptic stimuli at action potential frequencies up to 600 Hz (Wu and Kelly, 1993; Taschenberger and von Gersdorff, 2000). A number of Kþ channels have been shown to have specific functions in the auditory system that allow this degree of fidelity, particularly in the MNTB. These include KV1 channels, which contribute to a rapidly inactivating low-threshold Kþ current that suppresses the generation of multiple action potentials in response to synaptic currents (Brew and Forsythe, 1995; Brew et al., 2003), and KV3 channels, which contribute to a rapidly activating higher threshold Kþ current that decreases the duration of action potentials and allows high-frequency neuronal discharge locked to synaptic inputs (Wang et al., 1998; Macica et al., 2003; Song et al., 2005). Although the greatest evidence supports roles for KV1.1 and KV3.1 as mediators of these currents, there are likely contributions from other members of these voltage-gated Kþ channel subfamilies. A tonotopic gradient in Kþ currents has also been demonstrated for neurons of the MNTB, with medial (high-frequency responding) neurons demonstrating greater total KV1- and KV3-mediated Kþ currents than neurons in the lateral part of the nucleus (lower-frequency responding cells) (Li et al., 2001; von Hehn et al., 2004; Brew and Forsythe, 2005). There is evidence for additional Kþ conductance with different tonotopic variation as well, and recent evidence suggests that KNa channels play an important role in the fidelity of neuronal action potential timing at high frequencies. In fact, a major component of Kþ current in these cells is sensitive to cytoplasmic levels of Naþ (and Cl, see below), and increasing intracellular Naþ, as well as pharmacological manipulation directed to enhancement of KNa (see below), results in an increase in the accuracy of MNTB neuronal action potential timing (Yang et al., 2007).

7.3 CLONING AND IDENTIFICATION OF GENES ENCODING KNa CHANNELS: SLACK AND SLICK 7.3.1

The Slack Channel

Although recording from native KNa channels provided evidence of the importance of these channels in neuronal regulation, identification of the molecular substrate(s) of KNa was both elusive and important, since heterologous expression would allow for the more rapid and accurate discovery of pharmacological tools. In fact, KNa was one of the last remaining physiologically identified but molecularly uncharacterized macroscopic Kþ currents. In the late 1990s, near the end of the era of rapid cloning of the Kþ channel genome by “blast” homology searching of public DNA databases, a sequence designated as Slack (Slack is an acronym for sequence like a calcium-activated Kþ channel) was cloned from a rat cDNA library (Joiner et al., 1998). A related squence (f08b12.3) is present in the genome of Caenorhabditis elegans and this was later designated as C. elegans gene Slo2 (Yuan et al., 2000). The Slack sequence has regions

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FIGURE 7.1 The proposed structures of single Slack (a) and Slick (b) a subunits; explanations for each of the structural components are found in text. KNa channels are formed by four a subunits; currently it is unknown if a KNa channel tetramer can be formed by both Slack and Slick subunits, forming KNa channels with mixed properties.

of homology with Slo1, the gene encoding the BKCa channel protein (Butler et al., 1993; Dworetzky et al., 1994; McCobb et al., 1995; Gribkoff et al., 2001a). The overall Slack sequence corresponds to a Kþ channel a subunit, with a predicted full-length protein sequence of 1237 amino acids. Like most voltage-gated Kþ channels, it had a predicted structure consisting of six transmembrane domains and a putative signature GYG pore sequence (Fig. 7.1a). Unlike the Slo1/BKCa a subunit, it does not have the S0 transmembrane domain, which characterizes the unique structure of the largeconductance Ca2þ-activated Kþ channel and is the site for interaction with a number of BKCa b subunits (see Chapter 12 in this volume). Like the BKCa channel, it has a long C-terminal region. It also has a consensus PDZ-domain sequence in this long tail, suggesting it is capable of undergoing clustering. The protein has many sites for phosphorylation and, as will be described later, its activity is modulated by protein kinases (Santi et al., 2006). Interestingly, although in other respects it resembles a voltage-gated Kþ channel, it does not have the consensus voltage sensor sequence in S4 consisting of alternating positively charged residues. While Slack has little homology with Slo1 from the N-terminus to the pore, there are discrete regions of homology in the pore region and in parts of the long C-terminal tail. Overall, the homology of the Slack channel to Slo2/f08b12.3, Slo1 or the Slo3 channel of the testis is, however, only 7%. Despite this very low degree of homology, the discrete regions of homology are in critical regions, and the channel has been considered a member of the Slo gene family. Western blot analysis indicated that Slack is found in brain as 4.5 and 7.5 Kb transcripts and is widely distributed in brain. Specifically, in situ hybridization revealed very significant signal in the cortex, hippocampus, deep cerebellar nucleus, cerebellar Purkinje neurons, pons, preoptic nucleus, substantia nigra, and in a number of brainstem nuclei, including auditory nuclei such as the ventral cochlear nuclei and the MNTB. In a subsequent study, Bhattacharjee et al. (2005) used an affinity-purified antibody against the N-terminal region of rat Slack to evaluate the distribution of Slack in more detail. It is now known that this N-terminus is specific to one splice isoform of Slack, now termed Slack-B (Brown et al., 2006). They found that the

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FIGURE 7.2 Slack-B immunoreactivity in central neurons. (a). Prominent labeling is the substantia nigra (SNR) found in the subthalamic nuclei (STh). In the substantia nigra, staining is localized to neuronal cell bodies and axonal projections (b). Scale bars, 200 mm in (a), 100 mm in (b). (c) Slack-B immunolocalization in neurons of the frontal cerebral cortex (Bhattacharjee et al., 2002).

greatest expression, as reflected in the intensity of Slack-B immunoreactivity, was in the somata and axonal processes of neurons in the component nuclei of the brainstem trigeminal system and reticular formation. Intense staining was also observed in the vestibular nuclei, oculomotor nuclei, giant presynaptic terminals of the medial nucleus of the trapezoid body, deep cerebellar nuclei, substantia nigra, amygdala, and thalamus (Fig. 7.2a and b). In the cortex, significant staining was observed only in the frontal cortex (Fig. 7.2c). Slack-B protein is therefore widespread, but not ubiquitous. Expression of Slack channels in Xenopus oocytes and transiently transfected Chinese hamster ovary (CHO) cells produced an outwardly rectifying current under voltage–clamp that activated in both expression systems at potentials positive to 60 mV and had both instantaneous and slower activating components (Fig. 7.3) (Joiner et al., 1998). The single-channel conductance, recorded from inside-out

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FIGURE 7.3 Characteristics of Slack-B and Slick channels at the macroscopic and singlechannel levels. The Whole-cell currents in Slack-B or Slick-transfected CHO cells were recorded in response to 200 ms pulses to potentials between 120 and þ120 mV. Excised inside-out single-channel records are shown at a membrane potential of þ80 mV.

membrane patches bathed in symmetrical Kþ solution, was 40–65 pS with smaller subconductance states. Increased intracellular Ca2þ produced a significant inhibition of Slack channel activity recorded in both expression systems. In this initial study, the regions of sequence similarity with the Slo1 channel suggested that the Slack a subunit might form heteromultimers with Slo1 a subunits. To test this hypothesis, mRNA for both Slack and Slo1 was coinjected into Xenopus oocytes and the whole-cell currents and single-channel currents from inside-out membrane patches were examined. Following the coinjection, the pharmacology, kinetics, and single-channel conductance of Slack homomeric channels, Slo1 homomeric channels, and putative Slack/Slo1 heteromultimeric channels were analyzed. Currents from coinjected oocytes had kinetics and activation voltage ranges that were intermediate between Slack and Slo1. In addition, Slack channels were insensitive to the BKCa channel opener NS1619 and the BKCa channel blocker iberiotoxin, whereas the Slo1 channels were sensitive to both pharmacological agents. The currents resulting from coinjection of Slack/Slo1 were equally sensitive to NS1619, but at steady state were largely insensitive to iberiotoxin. These results were consistent with the formation of Slack/Slo1 heteromultimeric channels and inconsistent with the two independent populations of Slack and Slo1 in the coinjected oocytes. In the inside-out membrane patches, intermediate-conductance channels, intermediate between the single-channel conductance of either Slack (under these conditions in oocytes 25–65 pS) or Slo1 (
280 pS), were recorded with conductances between 80 and 190 pS. The channels in patches from coinjected oocytes were also activated by 1 mM Ca2þ applied to the intracellular face, similar to Slo1 channels but unlike Slack channels (which are inhibited by Ca2þ). This heteromultimerization phenomenon has also been observed with Slo2 channels from C. elegans, which are similar to Slack but like Slo1 channels are activated by Ca2þ and may form hybrid channels with Slo1 (Yuan et al., 2000, 2003). It therefore appears that the a subunits of these two channels may form channels with interesting mixed properties, both in their biophysical characteristics and pharmacology. Initially, it was thought that these results suggested that Slack may provide a substrate for the known diversity in native BKCa channels, but it is unknown whether this ever actually occurs in nature. Subsequent studies have provided

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evidence for another family member (Slick), a new alternative Slack sequence and an entirely different function for these channels as the substrate for KNa. 7.3.2

Identification of Slack as a Naþ-Activated Kþ Channel

In the initial studies following the cloning and expression of Slack Kþ channels, it was discovered that they were inhibited by [Ca2þ]in, but their sensitivity to [Naþ]in was not tested. The C. elegans ortholog, however, was found to be activated by both [Ca2þ]in and [Cl]in (Yuan et al., 2000). In 2003, a comprehensive study of both Slack and its C. elegans ortholog, Slo2, reported that Slack was activated by [Naþ]in and in the presence of elevated cytosolic Naþ, the currents closely resembled those of KNa (Yuan et al., 2003). In particular, the single-channel conductances were very similar to native KNa channels, varying significantly with different [Kþ]out and [Kþ]in. Although under the original recording conditions of Joiner et al. 1998, the rat Slack channels were reported to have single-channel conductance less that 100 pS, in this study the single-channel conductance in symmetrical 160 mM [Kþ]out/[Kþ]in was found to be 165 pS, similar to the conductance levels reported for native KNa channels (Yuan et al., 2003). The single-channel current amplitude also underwent considerable outward rectification at positive voltages. The Slack channels also exhibited multiple and prominent subconductance states, similar to native KNa channels. The whole-cell currents from Xenopus oocytes and the multichannel currents recorded from macropatches under physiological conditions were large outward currents (voltage steps between 80 and þ40 mV) with reversal potential near EK (78 mV in macropatches). The whole-cell currents did not demonstrate block at very positive voltages and had a slower activating component than was observed in macropatches. The C. elegans ortholog Slo2 was not activated by Naþ but, as reported earlier, was activated by Ca2þ. Both of these channels were also activated by [Cl]in. Although this had not been previously described for KNa channels, subsequent analyses have found Cl sensitivity of KNa channels in kidney and in MNTB neurons (Paulais et al., 2006; Yang et al., 2007). In the case of the C. elegans channel, the dependence on both [Ca2þ]in and [Cl]in is absolute, with no significant currents observed in the absence of either ion. In mammalian Slack, the dependence on [Cl]in is not absolute, and while it enhances channel activation, some degree of channel activation can be observed in Naþ alone. The effect of the two ions in Slack is cooperative or synergistic, in that the activation in the presence of both ions is greater than the sum of the activation by either ion alone. Channels in the Slo family, exemplified by the Slo1 BKCa channel, are known to possess regions in the long carboxy-terminal region that may underlie their sensitivity to ions. In the case of Slo1 BKCa channels, these consist of two RCK (regulator of Kþ conductance) domains, regions that have been previously shown in bacterial Ca2þ-activated Kþ channels to form a ring structure that responds to Ca2þ by regulating the opening and closing of the channel (Jiang et al., 2002; Parfenova et al., 2006), and a “Ca2þ bowl” believed to be the major site for the binding of Ca2þ (Schreiber and Salkoff, 1997). Slack channels also have two RCK domains (Fig. 7.1a and b) and a region homologous to the Ca2þ bowl; this latter region is highly

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homologous between Slo2 orthologs (Yuan et al., 2000, 2003). Ion sensing in this “bowl” region in Slo1 BKCa channels is accomplished by a series of negatively charged residues thought to coordinate Ca2þ, whereas in Slo2 channels, these residues are replaced by positively charged amino acids in comparable positions (Yuan et al., 2003). While these positive charges may help coordinate Cl, however, mutation of every charged residue of the bowl region of Slack to the neutral amino acid glutamine produces no change in the affinity of the channels to Cl or Naþ, indicating that this region is unlikely to control channel gating by these ions (Bhattacharjee et al., 2003). Slack has been shown to exist in at least two splice isoforms, which differ in their N-terminal domains (Brown et al., 2006). These two alternative splice variants are now known as Slack-B (the original cloned isoform) and Slack-A. As illustrated in Fig. 7.1a, the Slack-B has a longer N-terminus and results in slowly activating currents as described above, whereas the Slack-A has a shorter N-terminus and activates more rapidly, similar to Slick (see below). 7.3.3

The Slick Channel

In 2003, a second gene encoding a KNa channel was identified (Bhattacharjee et al., 2003). This channel now termed Slick (sequence like an intermediate-conductance Kþ channel) was found to be a rapidly gating Naþ- and Cl-sensitive Kþ channel with significant structural homology with Slack. Like Slack, Slick (Slo2.1) has six transmembrane domains and no apparent voltage-sensor region in S4, although, like Slack, the currents are voltage dependent. It also has a large C-terminal region with a highly conserved PDZ domain. Like Slack, Slick also has two RCK domains, and the overall homology of the channels is approximately 70% (Fig. 7.1b). The two channels (Slack-B versus Slick), however, differ significantly in several aspects, including faster activation kinetics for Slick channels than Slack-B (Fig. 7.3). Slack-B has an instantaneous component, but the majority of current is slower activating. However, as mentioned above, an alternative Slack transcript called Slack-A has now been described. Like Slick, Slack-A is more rapidly activating, and has a high degree of structural homology in its alternative N-terminal sequence with Slick’s N-terminal sequence. Unlike both Slack transcripts, Slick also has a higher sensitivity to [Cl]in, has some ability to conduct in the absence of high [Naþ]in, and, most important, has a consensus ATP binding motif in its C-terminal region. Slick is in fact an ATP-regulated channel. In the presence of 5 mM of MgATP in the recording pipettes, Slick current densities were reduced by as much as 80% at all tested voltages (Bhattacharjee et al., 2003). The effect of ATP was reproduced by slow or nonhydrolyzable analogues of ATP, and the effect was markedly reduced in the presence of ADP and abolished by mutation of the ATP binding motif. The inhibition of Slick currents by ATP was not significantly affected by the KATP channel inhibitor glybenclamide or the KATP channel opener diazoxide, suggesting that Slick represents a new type of ATP-mediated Kþ channel regulation. The distribution of Slick has been examined in the rat CNS using in situ and immunocytochemical techniques (Bhattacharjee et al., 2005). Very strong

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hybridization signals and immunoreactivity were observed in brainstem nuclei, including the MNTB, and the signal was seen in most of the areas where Slack is seen. These include the olfactory bulb, red nucleus, thalamus, facial nucleus, vestibular nucleus, pontine nucleus, oculomotor nucleus, deep cerebellar nuclei, and the substantia nigra. Most interesting, however, was the observation that Slick mRNA and immunoreactivity were found in some areas that do not express significant levels of Slack-B. These include hippocampal neurons in areas CA1, CA2, and CA3, the dentate gyrus, the supraoptic nucleus of the hypothalamus as well as other hypothalamic loci, and cortical neurons of layers II, III, and IV (Bhattacharjee et al., 2005). It is now known that Slack-B and Slick channels can form heteromultimers both in expression systems and in native neurons (Chen et al., 2006, 2007). It also appears, however, that Slick can operate independently as a KNa channel, given its solo expression in some structures. This pattern of overlapping and nonoverlapping distributions of expression of Slack and Slick can explain some or all of the observed heterogeneity in the KNa channels and currents in the literature, particularly when the alternative Slack-A transcript and possible Slack/Slick, heteromultimeric channels are factored in.

7.4 MODULATION OF KNa CHANNELS BY SECOND MESSENGERS Slack and Slick are differentially regulated by the activation of protein kinase C (Santi et al., 2006). The whole-cell Slack currents are increased by exposure of cells to the protein kinase C activators phorbol 12-myristate 13-acetate (PMA) or 1-oleoyl-2acetyl-sn-glycerol (OAG). In addition, the activation of protein kianse C slows the rate of activation of Slack currents. In contrast, the Slick currents are inhibited by the activation of protein kinase C. Each of these effects is prevented by inhibitors of protein kinase C. Similar modulation of Slack and Slick currents is observed in response to activation of Gaq-protein-coupled receptors (GqPCRs) that are coupled to activation of protein kinase C (Santi et al., 2006). When GqPCRs such as the M1 muscarinic cholinergic receptor or the mGluR1 metabotropic glutamate receptor are coexpressed with the KNa channel subunits in Xenopus oocytes, the application of receptor agonist strongly enhances Slack currents and inhibits Slick currents.

7.5 PHARMACOLOGY OF KNa CHANNELS The KNa channels are important regulators of neuronal excitability, and pharmacological agents that affect their activation can have great value as research tools and as potential therapeutic agents. As research tools, inhibitors can be used to pharmacologically dissect KNa currents from specific neurons, neuronal populations, or the whole animal, helping to determine the function of these channels in neuronal and behavioral regulation. Openers or activators of these channels can likewise be used to better understand the range and limits of KNa contributions to neuronal behavior. In the

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search for therapeutic agents that can modulate neuronal excitability (or cardiac excitability), including the response to insults such as ischemia, a well-developed pharmacology is a critical first step in the development of effective and specific drugs directed to this target. Inhibitors of specific Kþ channels have been discovered from a variety of sources over the last several decades, with specific peptide toxins providing the greatest potency and specificity. In the case of activators, which often have the most obvious therapeutic potential (by enhancing Kþ channel-mediated damping of cellular excitability), there was, however, only a single class of Kþ channel (KATP channels) for which activators were known prior to approximately 1994. We now know that many other Kþ channels can be activated (their open probability increased) by small molecules. These include BKCa/Slo1 channels and neuronal KV7 (KCNQ) channels (McKay et al., 1994; Gribkoff et al., 1996, 2001b; Cooper and Jan, 2003; Gribkoff, 2008), and new reviews of the pharmacology of both of these Kþ channel subfamilies are found in Chapters 15 and 16 of this volume. Importantly, it has been observed that openers of these different channels often (but not always) have effects on other Kþ channels. Whether this represents some common mechanism(s) of modulation, for which differently configured “receptors” exist on different channel types, is currently unknown, but this observation provides an avenue for the initial discovery of compounds affecting additional Kþ channels without the need for large-scale screening campaigns. Using small libraries of compounds enriched for Kþ channel modulators, it can prove relatively simple to discover effective initial lead compounds, when coupled with cloned channels in an appropriate expression system. Prior to the cloning of Slack and Slick channels, it had proven very difficult to discover and characterize pharmacological agents that modulate KNa channels. Using native KNa channels and currents, only two agents had previously been described to affect KNa, both producing KNa inhibition. In ventricular myocytes, the antarrhythmic drugs bepripil and quinidine (Fig. 7.4b and c), previously known to block or inhibit other ion channels, were shown to also inhibit KNa channels (Mori et al., 1998; Li et al., 1999). Following the cloning of the first KNa channels, Slack-B channels and currents were employed to determine if quinidine and bepridil also inhibited cloned KNa channels, and also to determine if the logic discussed above could be used to discover KNa activators. Using two electrode voltage clamp recording from Slack mRNAinjected Xenopus oocytes and whole-cell and excised patch-clamp recordings from a stable Slack-expressing human embryonic kidney (HEK) cell line, the effects of the two putative KNa inhibitors were directly determined (Yang et al., 2006). In both expression systems, bepridil produced significant levels of inhibition. In Xenopus oocytes, the EC50 for inhibition by bepridil was between 5 and 10 mM. The sensitivity to bepridil was apparently higher when applied to HEK cells; the estimated IC50 for inhibition of whole-cell currents was approximately 1 mM. The inhibition was readily apparent at the single-channel level as well, with marked inhibition when applied to outside-out or inside-out membrane patches, an effect that was independent of [Naþ]in, because Naþ levels were directly controlled in the excised patch experiments. The inhibition of Slack-B currents by quinidine was also studied in these expression systems. Using Slack currents expressed transiently in the CHO cells, the inhibition of Slack currents by quinidine had previously been shown using a high (1 mM)

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(a) OH

OH

Cl

S

Cl

Cl

Cl

(b)

N

CH3 O CH3

N

(c) H2C

HC

H

N H

HO H

OCH3

N

FIGURE 7.4 Structures of the KNa opener bithionol (a) and the KNa inhibitors, bepridil (b) and quinidine (c).

concentration. Using oocytes and a stable Slack-expressing cell line, it was shown that quinidine inhibited KNa currents in both expression systems, with EC50 values
100 mM in oocytes and
90 mM in Slack-expressing HEK cells (Yang et al., 2006). Similar to bepridil, quinidine produced inhibition of single-channel open probability when applied to either the intracellular or extracellular face of excised membrane patches. This study also, and for the first time, demonstrated that Slack-B channels and macroscopic currents could be enhanced by a small molecule. Bithionol, a bis-phenol antiparasitic compound (Knodell et al., 1972; Powell and Lampert, 1973) (Fig. 7.4a), was chosen for these experiments for two reasons: its previously demonstrated ability to activate another member of the Slo family, the BKCa channel

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(Boissard and Gribkoff, unpublished results), and its commercial availability. While essentially playing a hunch that the compound would also activate Slack channels, bithionol proved a good one, effectively enhancing currents in both expression systems. In Xenopus oocytes, the EC50 for activation by bithionol was
10 mM, while in Slack-expressing HEK cells, the enhancement of whole-cell currents had an EC50 of
0.7 mM. As with bepridil (but not quinidine), the sensitivity of whole-cell currents in Slack-expressing HEK cells was about 10X higher than the sensitivity observed in Slack-expressing oocytes for reasons that are not yet understood. The potency of bithionol in HEK cells (or for that matter in oocytes) expressing Slack is quite reasonable for a Kþ channel opener, suggesting that the structural motifs associated with the binding of this compound are worth pursuing further. These compounds certainly are not ideal as specific pharmacological agents and all are capable of influencing the activity of other ion channels over comparable ranges. Nevertheless, particularly with bepridil and bithionol, the inhibition and activation of Slack currents, respectively, in both systems has been sufficiently potent and certainly effective enough to use these compounds as tools in select experiments. Great care has to be taken in such situations, however, so that effects on other targets are not interpreted as an effect on KNa channels. The search for more specific and potent KNa modulators will continue, but these initial successes demonstrate that smallmolecule modulators exist and that proper screening efforts should be fruitful. It should also be noted that although these molecules, particularly the opener, have thus far been studied systematically only in using Slack-B expressing systems, they are also effective on native neuronal KNa channels (Yang et al., 2006). Nevertheless, it is not yet known if different KNa channels have identical, similar, or very different pharmacological profiles. It should prove interesting and more than of academic interest to determine the degree to which the pharmacology of the two known subtypes of KNa channels are different.

7.6 CONCLUSIONS KNa channels have been known since the 1980s to be ligand-gated neuronal Kþ channels capable of regulating the responsiveness of excitable cells by virtue of their activation following accumulation of [Naþ]in. Over the last several years, two mammalian Kþ channels, Slack and Slick, have been shown to be KNa channels, responsive to both [Naþ]in and [Cl]in, with the characteristics of slowly and more rapidly activating KNa channels, respectively. The discovery of Cl sensitivity in cloned KNa channels was surprising, but following the identification of Slack and Slick as KNa channels, examples of native Cl-sensitive native KNa channels have been found (Paulais et al., 2006; Yang et al., 2007). Currently, it cannot be said with absolute certainty that all KNa channels are encoded by Slo2 channels, but the distributions of Slack and Slick, the common effects of pharmacological agents such as bepridil and bithionol on native and cloned KNa channels, as well as the effects of other modulators such as ATP, strongly suggest that Slack and Slick are subunits comprising native KNa channels. The question of the precise makeup of KNa channels, in terms of whether

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heteromultimeric combinations of Slack and Slick form native channels, is as yet unresolved and will likely require more sophisticated pharmacological and molecular manipulations to make a final determination. The functions of KNa channels in neurons are probably numerous and are adapted specifically for each cell type in which they are expressed, but certain generalities can certainly be made. Neuronal KNa channels are involved in the control of action potential burst length by virtue of their production of summating, long-lasting AHPs following action potential trains. Since [Naþ]in will be the highest after highfrequency, prolonged trains of action potentials and high [Naþ]in is generally required for their activation, KNa channels play a prominent role in neurons wherein a combination of Kþ conductances allow periods of high-frequency discharge without significant Naþ channel inactivation and/or persistent Naþ current is induced during bursts of action potentials. KNa channels can also be activated by Naþ entry through Naþ-permeable neurotransmitter receptors such as AMPA receptors (Nanou and El, 2007). KNa is not only suited as a mechanism of terminating bursts accompanied by Naþ-dependent DAPs, but also plays a role in maintaining fidelity of high-frequency action potentials trains in auditory nuclei such as the MNTB. In this latter nucleus, bithionol, shown to be an activator of Slack-B, enhanced the native KNa current and produced an increase in the accuracy of action potential timing at high stimulation frequencies (Yang et al., 2007), providing additional evidence for native KNa channels being some combination of Slack and Slick subunits. Initial pharmacological discovery had focused on using the newly cloned Slack-B channels to investigate previous observations that some antiarrythmic compounds blocked native KNa channels in cardiac myocytes and to discover initial lead KNa openers. The results have been promising, with bepridil (and to a lesser extent quinidine) found to block Slack-B channels at reasonable potency, and the bis-phenol compound bithionol shown to open both Slack-B channels and to enhance native KNa currents in MNTB neurons. All of these agents are leads and while reasonably potent, they lack a high degree of specificity, and there is as yet little published data on their effects on Slack-A or Slick. In select experiments, however, where the contributions of other conductances can be accounted for or negated using other tools, they can be used to examine the contributions of native channels to neuronal function. The focus of the search for new compounds will be on potency and selectivity. Of course it will also be of interest to determine if there is significant therapeutic utility of openers or inhibitors of KNa channels. Openers in particular would seem to have potential utility as antiepileptic compounds as well as utility in acute neurodegenerative disorders, including stroke and other forms of neuronal ischemia and traumatic brain injury. There is significant evidence that KNa channels function in cardiac myocytes as part of the cellular response to ischemia, because the levels of [Naþ]in required for activation of the channels is seldom reached except during pathological conditions. However, even this notion may be somewhat speculative. Increasing knowledge of the structural basis for the modulation of KNa channels by phosphorylation or other influences, many of which could conceivably alter the sensitivity of KNa channels to [Naþ]in or the combination of [Naþ]in and [Cl]in, may suggest a more dynamic role for KNa channels in cardiac cells. Nevertheless, KNa channels are particularly suited to

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act as a buffer for ischemic depolarization and cellular compromise in cardiac myocytes and many neurons. The Slo2 channel of C. elegans has been directly shown to be important in providing resistance to ischemia (Yuan et al., 2003). In KNa channels where the Slick subunit is part of the molecular substrate of native KNa channels, its sensitivity to ATP (where a decrease in ATP should release the channel from inhibition) seems tailor-made for responding to ischemic conditions, where [Naþ]in levels rise accompanied by a decrease in ATP. It will be interesting to determine the degree to which openers of these channels can delay phenomena associated with neuronal ischemia, such as hypoxic/anoxic depolarization (Aitken et al., 1998; Muller, 2000). These channels are important components of the regulatory “governor” on neuronal excitability, and future experiments using newly discovered modulators would ultimately provide very useful additional information on the roles of KNa channels, their molecular makeup, and possibly important therapies for a variety of neuronal disorders.

REFERENCES Aitken PG, Tombaugh GC, Turner DA, Somjen GG, 1998. Similar propagation of SD and hypoxic SD-like depolarization in rat hippocampus recorded optically and electrically. J Neurophysiol 80: 1514–1521. Bertrand D, Bader CR, Berheim L, Haimann C, 1989. KNa, a sodium-activated potassium current. Pflugers Arch 414 (Suppl 1): S76–S79. Bezanilla F, 2005. The origin of subconductance levels in voltage-gated Kþ channels. J Gen Physiol 126: 83–86. Bhattacharjee A, Gan L, Kaczmarek LK, 2002. Localization of the slack potassium channel in the rat central nervous system. J Comp Neurol 454: 241–254. Bhattacharjee A, Joiner WJ, Wu M, Yang Y, Sigworth FJ, Kaczmarek LK, 2003. Slick (Slo2.1), a rapidly-gating sodium-activated potassium channel inhibited by ATP. J Neurosci 23: 11681–11691. Bhattacharjee A, von Hehn CA, Mei X, Kaczmarek LK, 2005. Localization of the Naþ-activated Kþ channel Slick in the rat central nervous system. J Comp Neurol 484: 80–92. Bond CT, Maylie J, Adelman JP, 1999. Small-conductance calcium-activated potassium channels. Ann N Y Acad Sci 868: 370–378. Brew HM, Forsythe ID, 1995. Two voltage-dependent Kþ conductances with complementary functions in postsynaptic integration at a central auditory synapse. J Neurosci 15: 8011– 8022. Brew HM, Forsythe ID, 2005. Systematic variation of potassium current amplitudes across the tonotopic axis of the rat medial nucleus of the trapezoid body. Hear Res 206: 116–132. Brew HM, Hallows JL, Tempel BL, 2003. Hyperexcitability and reduced low threshold potassium currents in auditory neurons of mice lacking the channel subunit Kv1.1. J Physiol 548: 1–20. Brown MR, Spilianakis CG, von Hehn CA, Flavell RA, Bhattacharjee A, Kaczmarek LK, 2006. Genetic regulation of Slack channels via alternative promoters. Soc Neurosci Abstr 627: 18.

REFERENCES

211

Butler A, Tsunoda S, McCobb DP, Wei A, Salkoff L, 1993. mSlo, a complex mouse gene encoding “maxi” calcium-activated potassium channels. Science 261: 221–224. Chapman ML, VanDongen AMJ, 2005. K channel subconductance levels result from heteromeric pore conformations. J Gen Physiol 126: 87–103. Chen H, Yan Y, Gazula VR, Yang Y, Ferreira G, Sigworth FJ, Kaczmarek LK, 2006. The formation and trafficking of heteromeric Slick/Slack sodium-activated potassium channels are determined by the Slack N-terminus. Soc Neurosci Abstr 627: 16. Chen H, Yan Y, Gazula VR, Yang Y, Ferreira G, Sigworth FJ, Salkoff L, Kaczmarek LK, 2007. PKC regulation of Slack/Slick heteromeric potassium channels is distinct from that of homomeric channels. Soc Neurosci Abstr 43: 13. Cooper EC, Jan LY, 2003. M-channels: neurological diseases, neuromodulation, and drug development. Arch Neurol 60: 496–500. Dani JA, Fox JA, 1991. Examination of subconductance levels arising from a single ion channel. J Theor Biol 153: 401–423. Descalzo VF, Nowak LG, Brumberg JC, McCormick DA, Sanchez-Vives MV, 2005. Slow adaptation in fast-spiking neurons of visual cortex. J Neurophysiol 93: 1111–1118. Dryer SE, 1994. Naþ-activated Kþ channels: a new family of large-conductance ion channels. Trends Neurosci 17: 155–160. Dworetzky SI, Trojnacki JT, Gribkoff VK, 1994. Cloning and expression of a human largeconductance calcium-activated potassium channel. Brain Res Mol Brain Res 27: 189–193. Dworetzky SI, Boissard CG, Lum-Ragan JT, McKay MC, Post-Munson DJ, Trojnacki JT, Chang CP, Gribkoff VK, 1996. Phenotypic alteration of a human BK (hSlo) channel by hSlo beta subunit coexpression: changes in blocker sensitivity, activation/relaxation and inactivation kinetics, and protein kinase A modulation. J Neurosci 16: 4543–4550. Egan TM, Dagan D, Kupper J, Levitan IB, 1992a. Naþ-activated Kþ channels are widely distributed in rat CNS and in Xenopus oocytes. Brain Res 584: 319–321. Egan TM, Dagan D, Kupper J, Levitan IB, 1992b Properties and rundown of sodium-activated potassium channels in rat olfactory bulb neurons. J Neurosci 12: 1964–1976. Faber ES, Sah P, 2005. Independent roles of calcium and voltage-dependent potassium currents in controlling spike frequency adaptation in lateral amygdala pyramidal neurons. Eur J Neurosci 22: 1627–1635. Fox JA, 1987. Ion channel subconductance states. J Membr Biol 97: 1–8. Franceschetti S, Buzio S, Sancini G, Panzica F, Avanzini G, 1993. Expression of intrinsic bursting properties in neurons of maturing sensorimotor cortex. Neurosci Lett 162: 25–28. Franceschetti S, Lavazza T, Curia G, Aracri P, Panzica F, Sancini G, Avanzini G, Magistretti J, 2003. Naþ-activated Kþ current contributes to postexcitatory hyperpolarization in neocortical intrinsically bursting neurons. J Neurophysiol 89: 2101–2111. Gean PW, Shinnick-Gallagher P, 1989. The transient potassium current, the A-current, is involved in spike frequency adaptation in rat amygdala neurons. Brain Res 480: 160–169. Gribkoff VK, 2008. The therapeutic potential of neuronal Kv7 (KCNQ) channel modulators: an update. Expert Opin Ther Targets 12: 565–581. Gribkoff VK, Lum-Ragan JT, Boissard CG, Post-Munson DJ, Meanwell NA, Starrett JE Jr, Kozlowski ES, Romine JL, Trojnacki JT, McKay MC, Zhong J, Dworetzky SI, 1996. Effects of channel modulators on cloned large-conductance calcium-activated potassium channels. Mol Pharmacol 50: 206–217.

212

REGULATION OF NEURONAL EXCITABILITY

Gribkoff VK, Starrett JE Jr, Dworetzky SI, 2001a. Maxi-K potassium channels: form, function, and modulation of a class of endogenous regulators of intracellular calcium. Neuroscientist 7: 166–177. Gribkoff VK, Starrett JE Jr, Dworetzky SI, Hewawasam P, Boissard CG, Cook DA, Frantz SW, Heman K, Hibbard JR, Huston K, Johnson G, Krishnan BS, Kinney GG, Lombardo LA, Meanwell NA, Molinoff PB, Myers RA, Moon SL, Ortiz A, Pajor L, Pieschl RL, Post-Munson DJ, Signor LJ, Srinivas N, Taber MT, Thalody G, Trojnacki JT, Wiener H, Yeleswaram K, Yeola SW, 2001b. Targeting acute ischemic stroke with a calcium-sensitive opener of maxi-K potassium channels. Nat Med 7: 471–477. Gu N, Vervaeke K, Storm JF, 2007. BK potassium channels facilitate high-frequency firing and cause early spike frequency adaptation in rat CA1 hippocampal pyramidal cells. J Physiol 580: 859–882. Haimann C, Bader CR, 1989. Sodium-activated potassium channel in avian sensory neurons. Cell Biol Int Rep 13: 1133–1139. Haimann C, Bernheim L, Bertrand D, Bader CR, 1990. Potassium current activated by intracellular sodium in quail trigeminal ganglion neurons. J Gen Physiol 95: 961– 979. Haimann C, Magistretti J, Pozzi B, 1992. Sodium-activated potassium current in sensory neurons: a comparison of cell-attached and cell-free single-channel activities. Pflugers Arch 422: 287–294. Hess D, Nanou E, El MA, 2007. Characterization of Naþ-activated Kþ currents in larval lamprey spinal cord neurons. J Neurophysiol 97: 3484–3493. Ishii TM, Silvia C, Hirschberg B, Bond CT, Adelman JP, Maylie J, 1997. A human intermediate conductance calcium-activated potassium channel. Proc Natl Acad Sci USA 94: 11651–11656. Istvan PJ, Zarzecki P, 1994. Intrinsic discharge patterns and somatosensory inputs for neurons in raccoon primary somatosensory cortex. J Neurophysiol 72: 2827–2839. Jiang Y, Lee A, Chen J, Cadene M, Chait BT, MacKinnon R, 2002. Crystal structure and mechanism of a calcium-gated potassium channel. Nature 417: 515–522. Joiner WJ, Wang LY, Tang MD, Kaczmarek LK, 1997. hSK4, a member of a novel subfamily of calcium-activated potassium channels. Proc Natl Acad Sci USA 94: 11013–11018. Joiner WJ, Tang MD, Wang LY, Dworetzky SI, Boissard CG, Gan L, Gribkoff VK, Kaczmarek LK, 1998. Formation of intermediate-conductance calcium-activated potassium channels by interaction of Slack and Slo subunits. Nat Neurosci 1: 462–469. Kameyama M, Kakei M, Sato R, Shibasaki T, Matsuda H, Irisawa H, 1984. Intracellular Naþ activates a Kþ channel in mammalian cardiac cells. Nature 309: 354–356. Khawaja FA, Alonso AA, Bourque CW, 2007. Ca2þ-dependent Kþ currents and spikefrequency adaptation in medial entorhinal cortex layer II stellate cells. Hippocampus 17: 1143–1148. Kim U, McCormick DA, 1998. Functional and ionic properties of a slow afterhyperpolarization in ferret perigeniculate neurons in vitro. J Neurophysiol 80: 1222–1235. Knodell RG, Kirsch E, Rygg GC, 1972. Fascioliasis. response to bithionol Calif Med 117: 72–74. Kohler M, Hirschberg B, Bond CT, Kinzie JM, Marrion NV, Maylie J, Adelman JP, 1996. Small-conductance, calcium-activated potassium channels from mammalian brain. Science 273: 1709–1714.

REFERENCES

213

Kubota M, Saito N, 1991. Sodium- and calcium-dependent conductances of neurones in the zebra finch hyperstriatum ventrale pars caudale in vitro. J Physiol 440: 131–142. Li W, Kaczmarek LK, Perney TM, 2001. Localization of two high threshold potassium channels in the rat central auditory system. J Comp Neurol 437: 196–218. Li Y, Sato T, Arita M, 1999. Bepridil blunts the shortening of action potential duration caused by metabolic inhibition via blockade of ATP-sensitive Kþ channels and Naþ-activated Kþ channels. J Pharmacol Exp Ther 291: 562–568. Liu X, Stan Leung L, 2004. Sodium-activated potassium conductance participates in the depolarizing afterpotential following a single action potential in rat hippocampal CA1 pyramidal cells. Brain Res 1023: 185–192. Luk HN, Carmeliet E, 1990. Naþ-activated Kþ current in cardiac cells: rectification, open probability, block and role in digitalis toxicity. Pflugers Arch 416: 766–768. Macica CM, von Hehn CA, Wang LY, Ho CS, Yokoyama S, Joho RH, Kaczmarek LK, 2003. Modulation of the Kv3.1b potassium channel isoform adjusts the fidelity of the firing pattern of auditory neurons. J Neurosci 23: 1133–1141. Martin AR, Dryer SE, 1989. Potassium channels activated by sodium. Q J Exp Physiol 74: 1033–1041. McCobb DP, Fowler NL, Featherstone T, Lingle CJ, Saito M, Krause JE, Salkoff L, 1995. A human calcium-activated potassium channel gene expressed in vascular smooth muscle. Am J Physiol 269: H767–H777. McKay MC, Dworetzky SI, Meanwell NA, Olesen SP, Reinhart PH, Levitan IB, Adelman JP, Gribkoff VK, 1994. Opening of large-conductance calcium-activated potassium channels by the substituted benzimidazolone NS004. J Neurophysiol 71: 1873–1882. Metherate R, Aramakis VB, 1999. Intrinsic electrophysiology of neurons in thalamorecipient layers of developing rat auditory cortex. Brain Res Dev Brain Res 115: 131–144. Mistry DK, Tripathi O, Chapman RA, 1996. The occurrence of stable subconductance levels in Naþ-activated Kþ channels in excised membrane patches from guinea-pig ventricular myocytes. Exp Physiol 81: 899–907. Mori K, Kobayashi S, Saito T, Masuda Y, Nakaya H, 1998. Inhibitory effects of class I and IV antiarrhythmic drugs on the Naþ-activated Kþ channel current in guinea pig ventricular cells. Naunyn Schmiedebergs Arch Pharmacol 358: 641–648. Moss GW, Moczydlowski E, 1996. Rectifying conductance substates in a large conductance Ca2þ-activated Kþ channel: evidence for a fluctuating barrier mechanism. J Gen Physiol 107: 47–68. Muller M, 2000. Effects of chloride transport inhibition and chloride substitution on neuron function and on hypoxic spreading-depression-like depolarization in rat hippocampal slices. Neuroscience 97: 33–45. Nanou E, El MA, 2007. A postsynaptic negative feedback mediated by coupling between AMPA receptors and Naþ-activated Kþ channels in spinal cord neurones. Eur J Neurosci 25: 445–450. Nedergaard S, 2004. A Ca2þ-independent slow afterhyperpolarization in substantia nigra compacta neurons. Neuroscience 125: 841–852. Niu XW, Meech RW, 2000. Potassium inhibition of sodium-activated potassium (KNa) channels in guinea-pig ventricular myocytes. J Physiol 526: 81–90. Parfenova LV, Crane BM, Rothberg BS, 2006. Modulation of MthK potassium channel activity at the intracellular entrance to the pore. J Biol Chem 281: 21131–21138.

214

REGULATION OF NEURONAL EXCITABILITY

Patlak JB, 1993. Measuring kinetics of complex single ion channel data using mean-variance histograms. Biophys J 65: 29–42. Paulais M, Lachheb S, Teulon J, 2006. A Naþ- and Cl -activated Kþ channel in the thick ascending limb of mouse kidney. J Gen Physiol 127: 205–215. Pedarzani P, McCutcheon JE, Rogge G, Jensen BS, Christophersen P, Hougaard C, Strobaek D, Stocker M, 2005. Specific enhancement of SK channel activity selectively potentiates the afterhyperpolarizing current IAHP and modulates the firing properties of hippocampal pyramidal neurons. J Biol Chem 280: 41404–41411. Powell HC, Lampert PW, 1973. Bithionol: a possible substitute for hexachlorophene. Pediatrics 52: 859–861. Rodrigo GC, 1993. The Naþ-dependence of Naþ-activated Kþ-channels (IKNa) in guinea pig ventricular myocytes is different in excised inside/out patches and cell attached patches. Pflugers Arch 422: 530–532. Rodrigo GC, Chapman RA, 1990. A sodium-activated potassium current in intact ventricular myocytes isolated from the guinea-pig heart. Exp Physiol 75: 839–842. Safronov BV, Vogel W, 1996. Properties and functions of Naþ-activated Kþ channels in the soma of rat motoneurones. J Physiol 497 (Part 3): 727–734. Salkoff L, Butler A, Ferreira G, Santi C, Wei A, 2006. High-conductance potassium channels of the SLO family. Nat Rev Neurosci 7: 921–931. Sanabria ER, Su H, Yaari Y, 2001. Initiation of network bursts by Ca2þ-dependent intrinsic bursting in the rat pilocarpine model of temporal lobe epilepsy. J Physiol 532: 205–216. Sanchez-Vives MV, Nowak LG, McCormick DA, 2000a. Cellular mechanisms of long-lasting adaptation in visual cortical neurons in vitro. J Neurosci 20: 4286–4299. Sanchez-Vives MV, Nowak LG, McCormick DA, 2000b. Membrane mechanisms underlying contrast adaptation in cat area 17 in vivo. J Neurosci 20: 4267–4285. Sanguinetti MC, 1990. Naþ 1-activated and ATP-sensitive Kþ channels in the heart. Prog Clin Biol Res 334: 85–109. Santi CM, Ferreira G, Yang B, Gazula VR, Butler A, Wei A, Kaczmarek LK, Salkoff L, 2006. Opposite regulation of Slick and Slack Kþ channels by neuromodulators. J Neurosci 26: 5059–5068. Schreiber M, Salkoff L, 1997. A novel calcium-sensing domain in the BK channel. Biophys J 73: 1355–1363. Schwindt PC, Spain WJ, Crill WE, 1989. Long-lasting reduction of excitability by a sodiumdependent potassium current in cat neocortical neurons. J Neurophysiol 61: 233–244. Silva-Barrat C, Araneda S, Menini C, Champagnat J, Naquet R, 1992. Burst generation in neocortical neurons after GABA withdrawal in the rat. J Neurophysiol 67: 715–727. Sipila ST, Huttu K, Voipio J, Kaila K, 2006. Intrinsic bursting of immature CA3 pyramidal neurons and consequent giant depolarizing potentials are driven by a persistent Naþ current and terminated by a slow Ca2þ-activated Kþ current. Eur J Neurosci 23: 2330–2338. Song P, Yang Y, Barnes-Davies M, Bhattacharjee A, Hamann M, Forsythe ID, Oliver DL, Kaczmarek LK, 2005. Acoustic environment determines phosphorylation state of the Kv3.1 potassium channel in auditory neurons. Nat Neurosci 8: 1335–1342. Taschenberger H, von GH, 2000. Fine-tuning an auditory synapse for speed and fidelity: developmental changes in presynaptic waveform, EPSC kinetics, and synaptic plasticity. J Neurosci 20: 9162–9173.

REFERENCES

215

Traub RD, Wong RK, 1982. Cellular mechanism of neuronal synchronization in epilepsy. Science 216: 745–747. von Hehn CAA, Bhattacharjee A, Kaczmarek LK, 2004. Loss of Kv3.1 tonotopicity and alterations in CREB signaling in central auditory neurons of hearing impaired mice. J Neurosci 24: 1936–1940. Veldkamp MW, Vereecke J, Carmeliet E, 1994. Effects of intracellular sodium and hydrogen ion on the sodium-activated potassium channel in isolated patches from guinea pig ventricular myocytes. Cardiovasc Res 28: 1036–1041. Wallen P, Robertson B, Cangiano L, Low P, Bhattacharjee A, Kaczmarek LK, Grillner S, 2007. Sodium-dependent potassium channels of a Slack-like subtype contribute to the slow afterhyperpolarization in lamprey spinal neurons. J Physiol 585: 75–90. Wang Z, Kimitsuki T, Noma A, 1991. Conductance properties of the Naþ-activated Kþ channel in guinea-pig ventricular cells. J Physiol 433: 241–257. Wang LY, Gan L, Forsythe ID, Kaczmarek LK, 1998. Contribution of the Kv3.1 potassium channel to high-frequency firing in mouse auditory neurons. J Physiol 509 (Part 1): 183–194. Wikstrom MA, El MA, 1998. Calcium influx through N- and P/Q-type channels activate apamin-sensitive calcium-dependent potassium channels generating the late afterhyperpolarization in lamprey spinal neurons. Eur J Neurosci 10: 1528–1532. Wu SH, Kelly JB, 1993. Response of neurons in the lateral superior olive and medial nucleus of the trapezoid body to repetitive stimulation: intracellular and extracellular recordings from mouse brain slice. Hear Res 68: 189–201. Yang B, Gribkoff VK, Pan J, Damagnez V, Dworetzky SI, Boissard CG, Bhattacharjee A, Yan Y, Sigworth FJ, Kaczmarek LK, 2006. Pharmacological activation and inhibition of Slack (Slo2.2) channels. Neuropharmacology 51: 896–906. Yang B, Desai R, Kaczmarek LK, 2007. Slack and Slick KNa channels regulate the accuracy of timing of auditory neurons. J Neurosci 27: 2617–2627. Yuan A, Dourado M, Butler A, Walton N, Wei A, Salkoff L, 2000. SLO-2, a Kþ channel with an unusual Cl- dependence. Nat Neurosci 3: 771–779. Yuan A, Santi CM, Wei A, Wang ZW, Pollak K, Nonet M, Kaczmarek L, Crowder CM, Salkoff L, 2003. The sodium-activated potassium channel is encoded by a member of the Slo gene family. Neuron 37: 765–773.

PART II MODULATORY MECHANISMS AND INFLUENCES ON NEURONAL VOLTAGE-GATED ION CHANNEL FUNCTION

8 ALTERNATIVE SPLICING OF NEURONAL Cav2 CALCIUM CHANNELS DIANE LIPSCOMBE1, SUMMER E. ALLEN1, ANNETTE C. GRAY1,2, SPIRO MARANGOUDAKIS1, AND JESICA RAINGO1 1 2

Department of Neuroscience, Brown University, Providence, RI 02912, USA Department of Biology, Brandeis University, Waltham, MA 02454, USA

8.1 ALTERNATIVE PRE-mRNA SPLICING 8.1.1

Why Use Alternative Pre-mRNA Splicing?

The nervous system relies on access to finely tuned receptors and ion channels to maintain normal signaling and to adjust output in response to changing external cues. Alternative pre-mRNA splicing, a major step in RNA processing, allows cells to generate multiple proteins with subtly distinct activities from a single gene. This high-resolution control of cellular function would be much harder to achieve if a single gene generated only a single protein (Schwarz et al., 1988; Lipscombe, 2005). Neurons can optimize ion channel activity by mixing and matching protein modules, building on a backbone structure comprised of essential (constitutive) exons (Chemin et al., 2001; Maniatis and Tasic, 2002). For example, cell-specific alternative pre-mRNA splicing near ion channel domains that regulate channel gating kinetics can tailor excitability and firing rates in individual neurons (Rosenblatt et al., 1997; Oberholtzer, 1999; Ramanathan et al., 1999; Baranauskas et al., 2003). By adding or subtracting a number of discrete protein modules, alternative pre-mRNA splicing could achieve incremental control over channel function and support smooth transitions between a Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

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continuum of channel activities (Schwarz et al., 1988). On the other hand, the inclusion of an exon might turn a signaling pathway on or off by creating or disrupting interactions with signaling proteins and molecules. In such cases, alternative splicing has been compared to a molecular on/off switch (Mu et al., 2003; Chaudhuri et al., 2004; Raingo et al., 2007). Alternative pre-mRNA splicing is well documented: cell-specific splicing of down syndrome cell adhesion molecule (DSCAM) controls axonal targeting and cell–cell recognition during development (Schmucker et al., 2000); activity-dependent splicing of NMDA receptor RNA promotes export of NMDA receptors from the endoplasmic reticulum and controls surface expression at the synapse (Mu et al., 2003); cell-specific splicing of slo in hair cells of the inner ear underlies changes in calciumactivated potassium channels that set the resonant frequency of individual electrically tuned hair cells (Black, 1998; Shipston, 2001); the shift in alternative pre-mRNA splicing of SNAP-25 during development drives the transition from immature to mature synapses in the nervous system (Bark et al., 2004); and cell-specific pre-mRNA splicing of CaV1.3 L-type calcium channels in medium spiny neurons allows interaction with Shank and promotes channel localization at glutamatergic cortico–striatal synapses (Olson et al., 2005). Evidence shows that alternative pre-mRNA splicing is used extensively in the mammalian brain (Dredge et al., 2001; Lipscombe, 2005; Licatalosi and Darnell, 2006; Sharma and Black, 2006; Ule and Darnell, 2006; Zipursky et al., 2006). Furthermore, studies that link a specific splice isoform to an identified population of neurons and to a specific cell function support the biological importance of this process (Kollmar et al., 1997; Grabowski and Black, 2001; Baranauskas, 2004; Bell et al., 2004; Vendel et al., 2006; Raingo et al., 2007). 8.1.2

Basic Steps

Basic steps and different forms of alternative pre-mRNA splicing have been well described elsewhere (Smith and Valcarcel, 2000; Maniatis and Tasic, 2002; Black, 2003). This review will focus primarily on two common types of alternatively spliced exons: cassette exons that are included or excluded from the fully processed mRNA and mutually exclusive exons that are inserted individually but not together in the same mRNA (Fig. 8.1). The spliceosome – a complex molecular machine that controls the splicing of both constitutive and alternative exons – produces the catalytic reactions that remove introns and join exons during splicing. Whether the spliceosome causes a particular alternative exon to be included or excluded depends on the presence, absence, and/or ratios of specific splicing factors that associate with the pre-mRNA to promote specific splicing patterns (Smith and Valcarcel, 2000; Maniatis and Tasic, 2002). In turn, the cellular levels of these splicing factors depend on features such as cell type, developmental stage, and neuronal activity (Black, 2003). Splicing factors include both enhancer and repressor proteins. Splicing enhancers can recruit the spliceosome to the site of a particular alternative exon, ensuring that this exon is included. In contrast, splicing repressors can occlude a particular alternative exon from the spliceosome, causing this exon to be skipped. These enhancer and repressor factors work in concert with other RNA binding proteins to set the levels of specific mRNA splice isoforms required to support cellular function (Smith and Valcarcel,

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FIGURE 8.1 Two patterns of alternative pre-mRNA splicing used to generate different isoforms of CaV2.2. (a)The pattern involves the alternate expression of a cassette exon that is either included or skipped by the spliceosome. (b)The pattern involves inclusion of only one of the pair of exons (mutually exclusive). Introns are depicted with straight solid lines and exons with boxes. Constitutive exons are colored white and alternative exons are gray. When alternatively spliced exons are included and translated, they can confer unique functional properties to the protein. In (a), the inclusion of a cassette exon adds a domain that creates a new protein binding site. In (b), inclusion of either exon produces a protein binding domain, but only the second exon favors interaction with the specific target protein shown.

2000; Maniatis and Tasic, 2002; Black, 2003). Some splicing factors are cell and/or tissue specific (Fig. 8.2). Expression of these splicing factors can result in a change in the ratio of included or skipped alternative exons and can sometimes result in the expression of protein isoforms unique to specific cells or tissues (Fig. 8.2). Several RNA binding proteins involved in alternative splicing of neuronal pre-mRNAs are known, including Nova-1/2, Fox-1/2, Hu/elav family of proteins, CELF, neural PTB, KSRP (KH-type splicing regulatory protein), and NAPOR (neuroblastoma apoptosisrelated RNA-binding protein) (Min et al., 1997; Jensen et al., 2000; Polydorides et al., 2000; Ladd et al., 2001; Lisbin et al., 2001; Zhang et al., 2002; Jin et al., 2003). 8.1.3

Cell-Specific Alternative Splicing of Ion Channel Pre-mRNAs

Like all proteins, ion channels are highly sensitive to even small perturbations in their structure or chemical environment. However, for highly sensitive electrophysiological recording methods can monitor and quantify the impact of these changes on ion channel activity. Thus, those of us interested in ion channels can more readily assess the functional impact of alternative splicing on protein function and, when coupled to

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FIGURE 8.2 A large number of proteins and molecules act in concert to regulate alternative pre-mRNA splicing. In this hypothetical model of mutually exclusive splicing, all neurons express a number of regulatory proteins that either repress or enhance inclusion of one of the exon pair. In (a), repressor activity slightly exceeds enhancer activity on exon 1. Consequently, the spliceosome shows slight preference for exon 2, and the fraction of mRNAs containing exon 2 therefore slightly exceeds mRNAs containing exon 1. In (b), a repressor protein that binds to the intron upstream of exon 1 is expressed at low levels in neurons of dorsal root ganglia. This results in the spliceosome favoring exon 1 inclusion, and therefore mRNAs containing exon 1 are increased relative to total. Here, circles denote splicing enhancers, triangles denote splicing repressors, and a star denotes a tissue-specific repressor that is downregulated in dorsal root ganglia. (See the color version of this figure in Color Plate section.)

single-cell analysis, start to understand the impact of these activities within the context of specific neuronal populations (Lipscombe, 2005). Studies of alternative splicing of Slo transcripts that encode calcium-activated potassium channels nicely illustrate this point. In the chick cochlea, the pattern of alternative splicing of Slo pre-mRNAs varies with the tuning of hair cells along the basilar membrane to create a tonotopic map (Rosenblatt et al., 1997; Black, 1998; Fettiplace and Fuchs, 1999; Oberholtzer, 1999; Ramanathan et al., 1999). The correlation between the expression of Slo splice isoforms and tuning frequency in chick hair cells suggests that alternative splicing optimizes neuronal excitability over a wide range of tonal frequencies (Black, 1998; Fettiplace and Fuchs, 1999). Single-cell reverse transcriptase-polymerase chain reaction (RT-PCR) and electrophysiology have also been used to link the potassium channel splice isoform KV3.4a to the fast spiking phenotype of four different populations of neurons in the rat central nervous system (Baranauskas et al., 2003). In our studies of CaV2.2, we showed that alternative splicing in individual rat nociceptors underlies the unusually high sensitivity of calcium channels in these cells to inhibition by GABA and opioids (Raingo et al., 2007).

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8.1.4 How Many Voltage-Gated Calcium Channels are There? How Many Does the Brain Need? Currently, we can’t give precise answers to either of these questions, but we do know the numbers are high based on several lines of reasoning. First, we have genomic evidence from CaVa1 genes that encode the functional and structural core of all voltage-gated calcium channels; they span 100–800 kb of human genome sequence and each contains at least 50 exons (Soldatov, 1994; Lipscombe et al., 2002; Lipscombe and Castiglioni, 2004). Therefore, CaVa1 genes have the capacity to generate thousands of unique splice isoforms, as is the case for other genes far smaller than CaVa1 (Missler and Sudhof, 1998; Schmucker and Flanagan, 2004). To visualize the extent of alternative splicing for a given CaVa1 gene, we mapped reported sites of alternative splicing onto 2 of 10 human CaVa1 genes, CaV2.1 and CaV2.2, which encode the cores of P-type and N-type calcium channels, respectively (Fig. 8.3). From our own studies of CaV2.2 mRNAs, we know that exons e18a, e24a, e31a, and e37a/ e37b are expressed in specific regions of the nervous system or in specific types of neurons. We also know that these alternatively spliced exons uniquely influence Ntype calcium channel function (Lin et al., 1997, 1999, 2004; Pan and Lipscombe, 2000; Bell et al., 2004; Thaler et al., 2004; Gray et al., 2007; Raingo et al., 2007). Second, we have mRNA evidence, as a large number of CaVa1 subunit mRNA variants have been and continue to be isolated from a variety of tissues and cell lines (Koch et al., 1990; Perez-Reyes et al., 1990; Hui et al., 1991; Snutch et al., 1991; Diebold et al., 1992; Schneider et al., 1994; Williams et al., 1994; Marubio et al., 1996; Lin et al., 1997; Takimoto et al., 1997; Ligon et al., 1998; Mittman et al., 1999a; Cribbs et al., 2000; Chemin et al., 2001; Soong et al., 2002; Emerick et al., 2006). Third, we have functional evidence for a great diversity of voltage-gated calcium channels in the nervous system. Only 10 genes encode CaVa1 subunits, the functional core of all voltage-gated calcium channels, in mammalian genomes. Greater structural diversity than this is needed to explain the wide spectrum of cellular processes controlled by voltage-gated calcium channels (Delmas et al., 2000; Lipscombe et al., 2002; Lipscombe and Castiglioni, 2004). Fourth, genes that encode auxiliary calcium channel subunits, including all four CaVb and all three CaVa2d genes, also undergo alternative pre-mRNA splicing (Castellano and Perez-Reyes, 1994; Yamada et al., 2001; Helton and Horne, 2002; Helton et al., 2002; Lipscombe et al., 2002; Lipscombe and Castiglioni, 2004; Vendel et al., 2006). For example, alternative pre-mRNA splicing in the N-terminal domain of CaVb subunits regulates binding of synaptotagmin I and LC2 domain of microtubule-associated protein 1A (Vendel et al., 2006). The structural changes generated by alternative splicing of CaVa1 pre-mRNAs can regulate a wide range of channel properties, including biophysics, surface density, targeting, posttranslational modification, and coupling to downstream signaling pathways including G-protein-coupled receptors. Cell-specific factors coordinate alternative splicing of pre-mRNAs, and activities of these factors are influenced by tissue type, cell type, stage of development, neuronal activity, and disease (Diebold et al., 1992; Gidh-Jain et al., 1995; Angelotti and Hofmann, 1996; Lin et al., 1997, 1999; Takimoto et al., 1997; Welling et al., 1997; Vigues et al., 1998, 1999, 2002;

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FIGURE 8.3 Sites of alternatively spliced exons mapped on CaV2.1 and CaV2.2 proteins and the CaV2.2 gene. (a) and (b) illustrate the tetrameric structure and the two-dimensional transmembrane topology of CaV2.1 and CaV2.2 proteins, respectively. Approximate locations of alternatively spliced exons are highlighted with triangles and are numbered. The four main structural domains (I, II, III, and IV) are labeled and the constitutively expressed exons encoding each domain shown with black lines. (c) illustrates the human CaV2.2 gene. Exons are denoted as solid boxes and introns as lines. Constitutive exons are colored black and alternatively spliced exons colored gray. The gene structure was constructed from analysis of human genome sequence contig NT_023929.14. Small arrows indicate the location of alternative polyadenylation sites. Exons are numbered according to the previously published gene structure (Lipscombe et al., 2002). Figure adapted from Lipscombe and Castiglioni (2004). With kind permission of Springer Science and Business Media (See the color version of this figure in Color Plate section.)

Zuhlke et al., 1998; Bourinet et al., 1999; Pan and Lipscombe, 2000; Yang et al., 2000; Kaneko et al., 2002; Soong et al., 2002; Bell et al., 2004; Chaudhuri et al., 2004; Liao et al., 2004, 2005; Graf et al., 2005; Shen et al., 2006; Tiwari et al., 2006; Zhong et al., 2006; Altier et al., 2007; Gray et al., 2007; Raingo et al., 2007; Tang et al., 2007).

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Below, we briefly review alternative splicing of CaVa1 pre-mRNAs and then focus on processing of CaV2 pre-mRNAs that encode presynaptic calcium channels. 8.1.5 Which CaVa1 Subunit Domains are Modified by Alternative Pre-mRNA Splicing? Alternative splice sites exist throughout the coding regions of CaVa1 genes. At first glance, alternative splice sites appear more prevalent in the 30 halves of CaVa1 genes, but some of this bias arises from the 30 bias of sequences derived from mRNAs isolated from tissues as well as sequences derived in silico from the EST database. This is especially true for large genes such as Cava1. Certain general features of CaVa1 alternative splicing are worth pointing out. (i) Regions most divergent among different CaVa1 subunits contain the largest number of alternatively spliced exons, most notably the intracellular loops connecting domains II and III and the C-termini (Fig. 8.3). These regions play a critical role in linking the channel to target proteins and signaling molecules (Fig. 8.4). With additional information, we think alternatively spliced exons will have a similar representation in the N-terminus. (ii) Sequence differences among CaVa1 mRNA isoforms arising from alternative splicing may involve only a few nucleotides that insert a short peptide sequence (Lin et al., 1997; Bourinet et al., 1999) or hundreds of nucleotides that, if deleted, result in a frameshift and premature termination of translation (Soong et al., 2002). (iii) Similarly, the functional consequences of alternative splicing of CaVa1 pre-mRNAs may be quite subtle, such as changing channel activation kinetics (Lin et al., 2004; Murbartian et al., 2004), or profound, resulting in truncated, nonfunctional proteins (Soong et al., 2002). There are also interesting examples of alternative splicing of CaVa1 pre-mRNAs influencing channel pharmacology (Welling et al., 1997; Zuhlke et al., 1998; Beam, 1999; Bourinet et al., 1999; Fearon et al., 2000; Jimenez et al., 2000; Krovetz et al., 2000; Lacinova et al., 2000; Yatani and Kamp, 2000; Helton et al., 2002; Kaneko et al., 2002; Raingo et al., 2007). (iv) Certain sites of alternative splicing are conserved among paralogs and orthologs and are a general feature of all CaVa1 genes (e.g., cassette exons encoding part of the IVS3–IVS4 linker of CaVa1 subunits) (Lin et al., 1999; Lipscombe and Castiglioni, 2004), while other sites of alternative pre-mRNA splicing only exist in closely related CaVa1 genes (e.g., e37a and e37b in CaV2 genes (Gray et al., 2007)).

8.2 ALTERNATIVE SPLICING IN THE CaV2 GENES CaV2.1, CaV2.2, and CaV2.3 genes encode the core subunit of high-voltage-activated P-type, N-type, and R-type calcium channels respectively. CaV2 channels work together to control transmitter release from neurons of the mammalian nervous system. Their sequences andgenestructures are highly homologous.ThehumanCaV2.1gene,referred to as CACNA1A, is located on chromosome 19 at 19p13.2-p13.1; CaV2.2, CACNA1B, is located on Chromosome 9 at 9q34; and CaV2.3, CACNA1E, is on Chromosome 1 at

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FIGURE 8.4 Proteins that interact with the Cav2.2 subunit mapped onto a two-dimensional model of the channel. Several proteins interact with Cav2.2 channels, most notably in the II–III intracellular linker and C-terminus. A number of proteins associated with the synaptic machinery, and with synaptic transmission in general, interact with the II–III linker of Cav2.2, including CSP (cysteine string protein) (Miller et al., 2003), SNAP-25 (Catterall, 1999), synaptotagmin 1 (Sheng et al., 1997), syntaxin 1A (Sheng et al., 1994), huntingtin (Swayne et al., 2005), RGS12 (Richman and Diverse-Pierluissi, 2004), and b-arrestin (Puckerin et al., 2006). The asterisk (*) denotes proteins whose binding is either known or expected to be modified by alternative splicing. Binding of SNAP-25, synaptotagmin, and syntaxin is isoform dependent (Catterall, 1999). A CaV2.2 splice isoform lacking most of the II–III linker (e19 through e21) (Kaneko et al., 2002) theoretically should not interact with proteins that bind to this region. A second group of proteins interacts with the C-terminus, including 14-3-3 (Li et al., 2006), CASK (Maximov and Bezprozvanny, 2002), ENH (enigma homologue protein) (Maeno-Hikichi et al., 2003), Gaq (Simen et al., 2001), Gbg (De Waard et al., 2005), Mint1 (Maximov et al., 1999), PP2Ca (Li et al., 2005), RIM binding protein (RBP) (Hibino et al., 2002), and Tctex1 (Lai et al., 2005). CASK and Mint1 do not bind to a truncated splice isoform of CaV2.2 (Maximov et al., 1999; Maximov and Bezprozvanny, 2002). Theoretically, RIM binding protein should not bind to the truncated splice isoform of CaV2.2 described by Bezprozvanny and colleagues (Maximov et al., 1999). Gbg and CaVb subunits interact with the I–II linker (De Waard et al., 2005). Based on their sites of interaction in the I–II loop, alternative splicing at e10 (described in Section 8.2.3) should not disrupt Gbg and Cavb binding directly. The N-terminus is not known to undergo alternative splicing, thus the Gbg interacting site (Canti et al., 1999) should also not be affected by splicing. (See the color version of this figure in Color Plate section.)

1q25-q31. Access to genome sequences from a number of species, including human, has greatly facilitated our ability to confirm sites of alternative splicing in these genes and to identify new sites. Splice isoforms of all three CaV2 genes have been identified and characterized to various extents (Hui et al., 1991; Schneider et al., 1994; Williams et al., 1994; Sakurai et al., 1995; Marubio et al., 1996; Klockner et al., 1997; Lin et al., 1997,

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1999, 2004; Takimoto et al., 1997; Welling et al., 1997; Zhuchenko et al., 1997; Kamphuis and Hendriksen, 1998; Ligon et al., 1998; Pereverzev et al., 1998, 2002; Perez-Reyes et al., 1998; Soldatov et al., 1998; Vigues et al., 1998, 1999, 2002; Zuhlke et al., 1998; Beam, 1999; Bourinetet al., 1999; Ghasemzadeh et al., 1999; Hanset al., 1999; Lu and Dunlap, 1999; Mittman et al., 1999a, 1999b; Cahill et al., 2000; Cribbs et al., 2000; Krovetz et al., 2000; Lacinova et al., 2000; Monteil et al., 2000; Pan and Lipscombe, 2000; Chemin et al., 2001; Lu et al., 2001; McRory et al., 2001; Raghib et al., 2001; Staes et al., 2001; Jagannathan et al., 2002; Kaneko et al., 2002; Maximov andBezprozvanny,2002;Soongetal.,2002;Tsunemi etal.,2002;Belletal.,2004;Harry et al., 2004; Liao et al., 2004, 2005; Tang et al., 2004, 2007; Thaler et al., 2004; Graf et al., 2005; Castiglioni et al., 2006; Emerick et al., 2006; Kanumilli et al., 2006; Khanna et al., 2006; Shen et al., 2006; Vendel et al., 2006; Altier et al., 2007; Chang et al., 2007; Ikeda and Dunlap, 2007; Raingo et al., 2007). Several splice isoforms of CaV2 genes give rise to functionally and, in some cases, pharmacologically distinct channels (Bourinet et al., 1999; Hans et al., 1999; Jimenez et al., 2000; Kaneko et al., 2002; Raingo et al., 2007). We review studies of CaV2 splice isoforms according to specific structural domains of CaVa1 subunits that contain sequence encoded by alternatively spliced exons. We start at the 30 end of the channel at the C-terminus, a region that we know most about, and work forward toward the 50 N-terminus (Figs. 8.3a and b). 8.2.1

The C-Terminus

The C-terminus of CaVa1 constitutes about one fourth of the channel protein, and likely regulates various aspects of calcium channel function (Fig. 8.4) (Wei et al., 1994; Maximov et al., 1999; Zuhlke et al., 1999; Gao et al., 2000; Hering et al., 2000; Ivanina et al., 2000; Kepplinger et al., 2000; Peterson et al., 2000; Simen et al., 2001; Staes et al., 2001; Maeno-Hikichi et al., 2003; Li et al., 2004; Chen et al., 2005, 2006; Lautermilch et al., 2005; Castiglioni et al., 2006; Raingo et al., 2007; Wykes et al., 2007). CaVa1 splice isoforms differing in their C-termini have distinct kinetic properties including inactivation, activation, and deactivation (CaV1.2, CaV2.1, CaV2.2, CaV3.3) (Soldatov et al., 1997; Krovetz et al., 2000; Restituito et al., 2000; Murbartian et al., 2004; Castiglioni et al., 2006), G-protein sensitivity (CaV2.2) (Simen et al., 2001; Raingo et al., 2007), subcellular targeting (CaV1.3, CaV2.1, CaV2.2) (Maximov et al., 1999; Maximov and Bezprozvanny, 2002; Olson et al., 2005; Khanna et al., 2006), current densities (CaV2.1, CaV2.2) (Soong et al., 2002; Castiglioni et al., 2006), and sensitivity to calcium calmodulin-dependent inactivation and facilitation (CaV1.3 and CaV2.1) (Chaudhuri et al., 2004; Shen et al., 2006; Calin-Jageman et al., 2007; Chang et al., 2007; Wykes et al., 2007). The C-terminus is the most divergent domain among CaVa1 genes and alternative splicing in this region/generates differently sized forms of CaVa1 subunits (Fig. 8.3) (Hui et al., 1991; Schneider et al., 1994; Williams et al., 1994; Klockner et al., 1997; Soldatov et al., 1997; Zhuchenko et al., 1997; Ligon et al., 1998; Bourinet et al., 1999; Lu and Dunlap, 1999; Maximov et al., 1999; Hering et al., 2000; Krovetz et al., 2000; McRory et al., 2001; Xu and Lipscombe, 2001; Pereverzev et al., 2002; Calin-Jageman et al., 2007). In CaV2.1, the

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C-terminus is encoded by exons 36–47 (e36–e37) (Ophoff et al., 1996), and e37a/ e37b, e43, e44, and e47 are subject to alternative splicing (Fig. 8.3) (Zhuchenko et al., 1997; Bourinet et al., 1999; Krovetz et al., 2000; Soong et al., 2002; Chaudhuri et al., 2004; Kanumilli et al., 2006; Chang et al., 2007; Chaudhuri et al., 2007). In CaV2.2, the C-terminus is encoded by e36–e46 (Lipscombe and Castiglioni, 2004), and two putative sites of alternative splicing involving e37a/e37b and e46 have been described (Fig. 8.3) (Kaneko et al., 2002; Bell et al., 2004; Castiglioni et al., 2006; Raingo et al., 2007). 8.2.1.1 Mutually Exclusive e37a and e37b Mutually exclusive e37a and e37b are found in all three CaV2 genes (Gray et al., 2007). The lengths of e37a and e37b are identical, each containing 97 nucleotides, and they differ by 9–14 amino acids with some variation in amino acid composition of both exons among the three CaV2 genes. The e37a/e37b splice sites of CaV2.1 and CaV2.2 have been studied in most detail. Expression patterns of CaV2.1e[37a] and CaV2.1e[37b] mRNAs (Bourinet et al., 1999; Chang et al., 2007) are distinct from the expression pattern of the equivalent splice isoforms of CaV2.2 (Bell et al., 2004), suggesting that this splice site might have evolved to serve different functions in N-type and P-type channels. In CaV2.1, nine amino acids differ between e37a and e37b and they encode different forms of a helix–loop–helix structural motif that may bind calcium (EF-hand) (Bourinet et al., 1999; Krovetz et al., 2000; Soong et al., 2002; Chaudhuri et al., 2004). Both isoforms are expressed in human and rat brains and in human spinal cord (Gray et al., 2007), and selection of e37a or e37b may be under developmental control (Chang et al., 2007). The activity of CaV2.1e[37a] isoforms can be augmented via a calcium-dependent mechanism called calcium-dependent facilitation, a phenomenon that is absent in CaV2.1e[37b] isoforms. A second site of alternative splicing in the C-terminus encoding region of CaV2.1 pre-mRNA that involves e46 also modifies the calcium-dependent facilitation of CaV2.1 channels. David Yue and colleagues have shown that combinatorial alternative splicing at these two sites in CaV2.1 pre-mRNAs can generate three different forms of the Ptype channel in which calcium-dependent facilitation can be completely on, completely off, or off but ready for activation if global intracellular calcium rises (Chaudhuri et al., 2004, 2007), thereby acting as a molecular switch with three settings. Cav2.2e[37a] is enriched in dorsal root ganglia. Exons equivalent to CaV2.1e [37a] and CaV2.1e[37b] are found in CaV2.2. Alternative splicing at these sites regulates current density and G-protein signaling of the N-type channel (Fig. 8.5). CaV2.2 mRNAs expressed in most neurons contain e37b, except in dorsal root ganglia nociceptors, which express both CaV2.2e[37a] and CaV2.2e[37b] mRNAs (Bell et al., 2004). Enrichment of e37a-containing CaV2.2 mRNAs in nociceptors points to a functional role for alternative splicing of e37a pre-mRNAs in nociception. This was recently confirmed in a collaborative study led by Gerald Zamponi (Altier et al., 2007). Using splice isoform-specific small interfering RNA, Zamponi and colleagues showed that e37a-containing N-type channels contribute to basal

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CaV2.2e[37b]

GPCR

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GPCR

AP

ICa

FIGURE 8.5 Hypothesized physiological impact of alternative pre-mRNA splicing at the e37a/e37b site in CaV2.2. Hypothetical calcium currents (ICa) induced by a train of action potentials (AP) in cells expressing CaV2.2e[37b] or CaV2.2 e[37a] channels. CaV2.2e[37a] channels express at higher density compared to CaV2.2 e[37b] (Bell et al., 2004; Castiglioni et al., 2006), but they are also more sensitive to inhibition following G-protein receptor activation (Raingo et al., 2007). G-protein receptor activation is indicated by shaded regions. CaV2.2e [37b] currents are inhibited following G-protein-coupled receptor (GPCR) activation but because inhibition is purely voltage dependent, it is relieved during repetitive stimulation. CaV2.2e[37a] currents are also inhibited by GPCR activation but because inhibition of this isoform involves both voltage-dependent and voltage-independent pathways, inhibition is greater when compared to CaV2.2e[37b] currents, and it is only partially relieved during repetitive stimulation (Raingo et al., 2007). This model proposes that neurons could augment N-type current density and their sensitivity to G-protein-coupled receptors by increasing the ratio of CaV2.2e[37a]/CaV2.2e[37b] mRNAs. (See the color version of this figure in Color Plate section.)

thermal nociception and to thermal and mechanical hyperalgesia associated with inflammatory and neuropathic pain (Altier et al., 2007). More generally, this study provides evidence that CaV2.2e[37a] channels are present at presynaptic terminals of primary afferents and that they regulate synaptic transmission. N-type calcium channel blockers administered intrathecally are powerful analgesics in a variety of animal pain models (Miljanich and Ramachandran, 1995; Bowersox et al., 1996; Brose et al., 1997; Altier and Zamponi, 2004; Miljanich, 2004; Bourinet and Zamponi, 2005), validating N-type channels as targets for treating neuropathic pain. Collectively, pharmacological and siRNA studies raise the possibility that an N-type channel blocker with higher potency on CaV2.2e[37a] than CaV2.2e[37b] could have analgesic properties and fewer side effects than current therapeutics. 8.2.1.2 Alternative Splicing Controls G-Protein Signaling and Sets the Gain of the N-Type Channel What is the biological significance of e37a inclusion in CaV2.2 channels in a subset of nociceptors? We think alternative splicing of e37a and e37b in CaV2.2 pre-mRNAs is used to regulate the sensitivity of N-type channels to G protein-mediated inhibition, which in turn sets the sensitivity of nociception to neurotransmitters and drugs that act through G-protein-coupled receptors (Hille et al., 1995; Ikeda and Dunlap, 1999) (Fig. 8.5). We found that voltage-independent

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inhibition mediated by Gi/o can only gain access to the N-type channel if the CaV2.2 subunit contains e37a (Raingo et al., 2007). Combined with our previous studies that show enrichment of e37a in nociceptors (Bell et al., 2004), our data point to a molecular basis for the high susceptibility of N-type currents in sensory neurons to voltageindependent inhibition following G-protein activation (Diverse-Pierluissi et al., 1997; Ikeda and Dunlap, 2007). In addition to a higher sensitivity to G-protein-coupled receptors, CaV2.2e[37a] splice isoforms support larger N-type currents in nociceptive neurons, as compared to the ubiquitous CaV2.2e[37b] isoforms (Bell et al., 2004; Castiglioni et al., 2006). Larger currents associated with e37a-containing channels impacts the amount of calcium entering cells in response to a variety of action potential waveforms (Castiglioni et al., 2006). Overall, our studies imply that e37a effectively increases the gain of N-type channels; it supports larger N-type currents but at the same time renders them more sensitive to inhibition by G-protein-coupled receptor activation (Fig. 8.5). 8.2.1.3 Voltage-Independent Inhibition As discussed above, e37a controls Gi/o access to the N-type channel and consequently controls the level of voltageindependent inhibition. A Gi/o-dependent form of voltage-independent inhibition of N-type currents is well described in chick sensory neurons (Diverse-Pierluissi et al., 1997), consistent with enrichment of e37a in sensory neurons. Functionally, inhibition of calcium channels via this Gi/o-dependent, voltage-independent pathway is expected to be insensitive to changes in the membrane potential in contrast to the most common form of G-protein-mediated inhibition of N-type currents that is voltage dependent and mediated by G-protein Gbg dimers. Consequently, N-type currents activated by brief stimuli are well inhibited via this voltage-dependent pathway. However, with intense, high-frequency stimuli, this inhibition of N-type currents is relieved. Thus, voltage-dependent inhibition acts like a high pass filter that attenuates low-frequency signals, but has less of an effect on high-frequency signals (Ikeda and Dunlap, 1999, 2007; Elmslie, 2003). In addition to Gi/o, Gq can also inhibit N-type currents via a voltage-independent mechanism. Gq-dependent inhibition of N-type currents is well described in sympathetic neurons (Hille et al., 1995; Ikeda and Dunlap, 1999; Kammermeier et al., 2000; Elmslie, 2003) and may depend on the expression of alternatively spliced exons in CaV2.2 other than e37a. 8.2.1.4 Other Sites of Alternative Pre-mRNA Splicing in the C-Terminus In CaV2.1, alternative splicing of exon 44 regulates channel inactivation kinetics of the human spinal cord P-type channel (Krovetz et al., 2000). In this same region of the CaV2.1 gene, others have found evidence from analyses of mRNA in human brain that e43 is also subject to alternative splicing, and that together with e44, this site modifies the efficiency of P-type channel expression (Soong et al., 2002). An alternate 30 acceptor site in the intron upstream of e47 of CaV2.1 introduces a frameshift and early truncation, which deletes e47 from the C-terminus, producing a short CaV2.1 isoform. E47 is large, encoding >230 amino acids in the longer splice isoform of CaV2.1. The two C-terminus isoforms of CaV2.1 are differentially expressed in human brain (Soong et al., 2002), spinal cord (Krovetz et al., 2000), and neuroblastoma

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cells (Hans et al., 1999). An interesting consequence of e47 truncation is that the shorter form of CaV2.1 cannot contribute to spinocerebellar ataxia type-6 because the SCA6 polyglutamine expansion that underlies this disorder is located in e47 (Zhuchenko et al., 1997). Some investigators have speculated that the observed upregulation of e47 inclusion during late development might help explain why spinocerebellar ataxia phenotype has a delayed onset (Soong et al., 2002). The other well-documented functional role of the C-termini of voltage-gated calcium channel CaVa1 subunits is subcellular targeting. Splice isoform-specific interactions between two scaffold proteins, Mint1 and CASK, and the C-termini of both CaV2.1 and CaV2.2 subunits have been reported (Maximov et al., 1999) (Fig. 8.4). Mint1 and CASK only interact with the long C-terminal splice isoforms that contain e47, suggesting that alternative splicing at this site might play a role in regulating the density of calcium channels at presynaptic nerve terminals (Maximov et al., 1999). E46 in the C-terminus of CaV2.2 is equivalent to e47 of CaV2.1; it is large (>3000 nucleotides in length), it contains a putative alternative 30 acceptor site that is hypothesized to generate two differently sized forms of CaV2.2 (Williams et al., 1992; Lu and Dunlap, 1999; Maximov and Bezprozvanny, 2002), and it has three sites of polyadenylation, of which two are used to generate differently sized CaV2.2 mRNAs in the nervous system (Schorge et al., 1999). The relative abundance of the short and long forms of CaV2.2 is not known, but functional studies suggest that they localize to different subcellular compartments in neurons. The long form of CaV2.2 targets to synapses via its interaction with Mint1 and CASK, two proteins involved in subcellular targeting, and the shorter isoform of CaV2.2 predominantly localizes to soma and dendrites (Maximov and Bezprozvanny, 2002). The function of e46 beyond regulating CaV2.2 targeting needs to be investigated. 8.2.2

A Short Linker on the Outside of the Channel

The extracellular linker in the forth domain (IV) that connects the third and fourth transmembrane spanning segments (IVS3–IVS4) of most voltage-gated calcium channels contains an exon that is alternatively expressed. Alternative splicing within the IVS3–IVS4 linker is a feature of CaVa1 pre-mRNAs of all mammals studied to date, including human as well as Drosophila (Lipscombe et al., 2002). The IVS3– IVS4 region of CaV channels can therefore accommodate sequence modification (Lipscombe et al., 2002). In CaV2 genes, an alternatively spliced cassette exon (e31a) encodes part of the IVS–IVS4 linker. E31a of the CaV2.1 and CaV2.2 genes only encode two amino acids, but the length and amino acid composition of e31a of the different CaVa1 genes vary. Where function has been analyzed, e31a impacts the time course and voltage dependence of calcium channel activation (Perez-Reyes et al., 1990; Snutch et al., 1991; Starr et al., 1991; Diebold et al., 1992; Barry et al., 1995; Ihara et al., 1995; Lin et al., 1997, 1999, 2004; Ligon et al., 1998; Bourinet et al., 1999; Hans et al., 1999; Krovetz et al., 2000). E31a of CaV2.1 encodes the dipeptide sequence asparagine–proline (NP). The NP encoding exon of the human CaV2.1 gene is located within approximately 9 kb of intron sequence and its inclusion in CaV2.1 mRNAs in the mammalian nervous system

232

ALTERNATIVE SPLICING OF NEURONAL Cav2 CALCIUM CHANNELS

is under regional control (Lin et al., 1997; Bourinet et al., 1999; Krovetz et al., 2000; Soong et al., 2002). The amino acids encoded by e31a in CaV2.1 are reported in some studies to slow both activation and inactivation kinetics of P-type currents (Hans et al., 1999; Krovetz et al., 2000) while others report no effect of E31a on P-type channel inactivation (Bourinet et al., 1999). However, researchers agree that the dipeptide NP encoded by E31a in CaV2.1 modulates w-agatoxin IVA sensitivity (Bourinet et al., 1999; Hans et al., 1999). E31a inclusion decreases toxin affinity 11-fold compared to CaV2.1 channels that lack the NP dipeptide. Alternative splicing of e31a in CaV2.1 premRNA may underlie some of the pharmacological differences among P-type currents in different neurons that are often referred to as P-type and Q-type currents (Sather et al., 1993; Stea et al., 1994; Randall and Tsien, 1995; Bourinet et al., 1999). The absence of e31a-containing CaV2.1 mRNAs in single Purkinje cells that express classic P-type currents with high sensitivity to w-agatoxin IVA is consistent with this hypothesis. A homologous 6 nucleotide e31a in CaV2.2 encodes glutamate–threonine (ET). E31a of CaV2.2 has been located in human, mouse, and rat genes. This exon is expressed in a tissue-specific pattern, and its presence alters N-type channel function. In rat and human, e31a-containing CaV2.2 mRNAs are preferentially expressed in neurons of the peripheral nervous system and are virtually absent from the central nervous system. Analysis of the human CaV2.2 gene reveals that e31a is contained within 14 kb of intron sequence and flanked by consensus ag–gt splice sites (Lin et al., 1997, 1999). E31a slows N-type channel activation and deactivation kinetics and shifts the voltage dependence of channel activation to slightly more depolarized voltages. Although e31a of CaV2.1 and CaV2.2 encode different dipeptides (NP and ET, respectively), they slow channel activation kinetics to a similar degree (Lin et al., 1999), consistent with the proximity of IVS3–IVS4 to the putative voltage sensor (IVS4). E31a appears to directly affect the putative voltage sensor of CaV2.2, based on our own analyses of gating currents (Lin et al., 2004). The steady-state voltage dependence of charge movement associated with N-type channel gating currents was not consistently different when e31a was present. However, “on” gating currents that precede channel opening decayed more slowly in CaV2.2 channels containing e31a when compared with gating currents measured from channels lacking e31a. A slower gating current is thought to correspond to slower movement of the charge sensor in the channel. Thus, alternative splicing in the extracellular IVS3–IVS4 linker of CaV2.2 affects the kinetics, but not the voltage dependence of N-type channel gating (Lin et al., 2004). Extracellular domains of presynaptic calcium channels have also been implicated in synapse stabilization (Nishimune et al., 2004). Postsynaptic b2 laminins bind directly to presynaptic CaV2.1 and CaV2.2 channels, thereby physically bridging the two membranes and drawing calcium channels into the synaptic cleft. This interaction between presynaptic calcium channels and postsynaptic laminins may be important for organizing neurotransmitter release sites in the active zone. It would be interesting to know if alternative splicing in extracellular S3–S4 linkers of domains III and IV influence b2 laminin binding to CaV2 channels. A positive

ALTERNATIVE SPLICING IN THE Cav2 GENES

233

finding could point to a role of e31a in synapse stabilization and might hint at why this exon is only found in N-type channels of peripheral and not central neurons. 8.2.3

The Intracellular Linker LII–III Connects Domains II and III

The intracellular linker LII–III in CaV2 channels binds presynaptic proteins and is essential for excitation–secretion coupling (Fig. 8.4). This region is encoded by e18 through e21 (Fig. 8.3) and in CaV2, LII–III connects the channel to sites of synaptic vesicle fusion by binding SNARE (soluble N-ethyl-maleimide-sensitive fusion attachment protein receptor) proteins (Catterall, 1999). LII–III also plays a role in targeting CaV2 channels to presynaptic nerve terminals (Leveque et al., 1994; Sheng et al., 1994; Mochida et al., 1996, 2003; Rettig et al., 1996, 1997; Tobi et al., 1998; Catterall, 1999; Harkins et al., 2004) and it influences the biophysical properties of CaV2 channels, particularly cumulative inactivation (Catterall, 1999; Degtiar et al., 2000; Thaler et al., 2004). CaV2 isoforms distinguished by amino acid sequence differences in their LII–III linkers are thus likely to have distinct functions. 8.2.3.1 Alternative Splicing of e18a in CaV2.2 and CaV2.3 The best studied site of alternative pre-mRNA splicing in the LII–III regions of CaV2.2 and CaV2.3 genes involves an alternatively spliced cassette e18a that lies between constitutively expressed e18 and e19. The CaV2.1 gene lacks an equivalent alternatively spliced exon in this region. Cassette e18a is found in all mammalian and chick CaV2.2 and CaV2.3 genes, implying that the sequences it encodes regulate important cellular processes (Gray et al., 2007). Indeed, the fugu CaV2.2 gene also contains a cassette e18a. In rat and human CaV2.2 genes, e18a is located within a
9 kb long genomic sequence flanked by consensus ag–gt dinucleotide splicing recognition sequences (Gray et al., 2007). The 30 intron also contains alternative dinucleotide splice acceptors, ag, separated by three nucleotides that encode the amino acid arginine (R758). We know that both splice junctions are utilized because CaV2.2 mRNAs that lack and contain this amino acid have been isolated (Coppola et al., 1994; Cahill et al., 2000; Pan and Lipscombe, 2000; Kaneko et al., 2002). However, we do not know the function of R758 in CaV2.2. E18a in CaV2.2 is 63 nucleotides long and encodes 21 amino acids. Inclusion of e18a in CaV2.2 mRNAs varies by region in the human nervous system; e18a is expressed at higher levels in spinal cord compared to whole brain (Gray et al., 2007). Similarly, in rat, we have shown that CaV2.2e[18a] mRNAs are especially prevalent in adult sympathetic ganglia (>80%), but are expressed at lower levels in rostral brain structures such as neocortex (80%. To summarize this section, the activity of M-channels in sympathetic neurons is tightly governed by plasma membrane PIP2 abundance, and PLC-induced PIP2 depletion underlies muscarinic and angiotensin II suppression of these channels. In contrast, bradykinin and P2Y receptor stimulation depress M-current via [Ca2þ]i signals. Why should bradykinin or P2Y receptors use another route to inhibit M-channels? We will try to answer this question in section 11.3.6 of this chapter. 11.2.4

Other Modes of Modulation

Other signaling pathways affect M-current activity in SCG neurons either directly or indirectly. Thus, different M-channel subunits are subject to multiple phosphorylations that either directly affect channel gating or change its sensitivity to GPCR modulation. Thus, Kv7.3, 7.4, and 7.5 are phosphorylated by Src (Gamper et al., 2003; Li et al., 2004b) or receptor tyrosine kinases (such as the EGF receptor) (Jia et al., 2007) at two tyrosines equivalent to Y67 (N-terminus) and Y349 (C-terminus) in Kv7.3 (Li et al., 2004b; Jia et al., 2007), an effect resulting in the decrease of current amplitude and slowing of kinetics. In addition, Kv7.2 can be phosphorylated at S541 and S534 in the channel C-terminus by PKC, which is recruited to the channel by the A-kinase anchoring protein AKAP79/150 (Hoshi et al., 2003). These phosphorylations result in an increase of channel sensitivity to muscarinic, but not bradykinin, inhibition (Hoshi et al., 2003, 2005), presumably by decreasing the affinity of PIP2 for the channel (similar to suggested mechanism of action of Ca2þ/CaM; Delmas and Brown, 2005). Consistent with this, muscarinic depression of M-current in AKAP150 knockout mice was substantially attenuated (Hoshi et al., 2005). Finally, M-channels can be inhibited by cyclic ADP-ribose (Bowden et al., 1999; Higashida et al., 2000) and augmented by physiological concentrations of H2O2 (Gamper et al., 2006), and M-current is somewhat pH sensitive (Prole et al., 2003), although it is not clear if any of these M-channel properties are used in intracellular signaling pathways.

300

REGULATION OF NEURONAL ION CHANNELS

11.3 MODULATION OF VOLTAGE-GATED Ca2þ CHANNELS 11.3.1

Overview

Voltage-gated Ca2þ channels (VGCCs) form a large family of Ca2þ-selective ion channels that are expressed in all types of excitable cells and mediate release of neurotransmitters from synaptic terminals, secretion of neuromediators and hormones by neurons and neuroendocrine cells, excitation–contraction coupling, and Ca2þ-dependent gene expression (Catterall, 2000). There are five types of VGCC currents (ICa): high-voltage-activated L-, N-, P/Q-, and R-types, and low-voltage-activated T-type. The pore-forming subunit of VGCC, called a1, has 24 transmembrane domains (TMD) organized in four 6-TMD repeats. a1 Subunits are coded by three gene subfamilies: Cav1–3. Cav1.1–1.4 code for the L-type channels; Cav2.1 for P/Q-, Cav2.2 for N-, Cav2.3 for R-, and Cav3.1–3.3 for T-type channels, respectively. VGCC are usually assembled with auxiliary subunits: b, a2d, and, in some cases, g. In rat SCG cells, ICa is >80% of the w-conotoxin GVIA-sensitive N-type (Plummer et al., 1989), but in murine SCG, ICa is a mixture of about 50% N-type, 20% L-type, 25% P/Q-type, and the rest R-type (Shapiro et al., 1999). The N-type channels in particular have been shown to be expressed in a number of alternatively spliced forms (see Chapter 8; Lin et al., 1999), perhaps accounting for some of the variability in the pharmacological profile and modulatory sensitivities of Cav2.2 channels in the nervous system.

11.3.2

A Fast and Direct Modulatory Pathway

The early work of Dunlap and Fischbach 1978 showed that several neurotransmitters, including norepinephrine (NE), 5-HT, and GABA could trigger depressions of ICa in chick ganglia via G-protein actions (Holz et al., 1986). In rat SCG neurons, a similar and rapid depression of ICa by muscarinic (Wanke et al., 1987), somatostatin (Ikeda and Schofield, 1989), adrenergic (Plummer et al., 1991), and peptidergic (Foucart et al., 1993) receptor stimulation was observed. This action is mediated by pertussistoxin (PTX)-sensitive (Go/i) G-proteins. This type of signal was shown to result from a direct action of G-protein (Forscher et al., 1986; Lipscombe et al., 1989; Bernheim et al., 1991; Shapiro and Hille, 1993) with the channels, resulting in a shift in the voltage dependence of activation (Marchetti et al., 1986; Ikeda et al., 1987). The biophysics was nicely analyzed by Bruce Bean, who described the G-protein unbound and bound channels as “willing” and “reluctant,” respectively, such that “reluctant” channels only open at much more depolarized potentials than do “willing” ones (Bean, 1989). The “willing/reluctant” model correctly predicts that G-protein binding itself to the channel should be voltage dependent (favored at hyperpolarized potentials, disfavored at depolarized ones), such that upon a physiological depolarization, only the (unbound) “willing” channels can initially open, but a maintained depolarization will induce unbinding of G-protein, conversion of “reluctant” channels to “willing” ones, and a seeming biphasic or “slowed” activation time course (Boland and Bean, 1993). Indeed, as a quick and easy assay of Ca2þ channel modulation, many investigators

MODULATION OF VOLTAGE-GATED Ca2þ CHANNELS

301

simply look for this signature of slowed activation. As for Go/i activation of GIRK inward rectifier channels (Logothetis et al., 1987), it was soon realized that the bg dimer of the G-protein was responsible for this direct action on the Ca2þ channel as well (Herlitze et al., 1996; Ikeda, 1996; Delmas et al., 1998b). A number of other labs also contemporaneously explored this issue, using other neurons or receptor types (Gross et al., 1989; Kasai and Aosaki, 1989; Bley and Tsien, 1990; Elmslie et al., 1990; Schofield, 1990; Penington et al., 1991; Scholz and Miller, 1991; Ikeda, 1992; Golard and Siegelbaum, 1993). The topic has also been nicely reviewed (Hille, 1994; Ikeda and Dunlap, 1999). 11.3.3

A Slower and Indirect Pathway

The use of other G-protein mechanisms of modulation of these channels, however, soon became clear. The best example remains that of muscarinic agonists in SCG cells, for which the same agonist initiates two distinct signals acting on the same channels, but via different receptor subtypes and modes of action. The first uses the M2 or M4 subtypes of receptors (Bernheim et al., 1992; Shapiro et al., 1999), PTXsensitive Go/i G-proteins, and the voltage-dependent “willling/reluctant” mechanism discussed above (Beech et al., 1992). Due to its direct nature, its top speed is very fast; at Nav 1.7 > Nav1.2

Nav1.5 > Nav1.7–Nav 1.2 > TTX-R

Nonselective

Selectivity (Most to Least Potent)

25

9

Yes

UseDependent Ratio

SNL, CCI, CFA

Formalin (phases 1 and 2)

Formalin (phases 1 and 2), CFA, CCI

Pain Models (In Vivo)

371

11

BA 42730A

Crobenetine (BIII 890 CL)

13B

13,

OH

N

OCF3

N

OH

O

O

O

N

S

OCH3

OCH3

O

NH

N

O

N

S

Cl

(Liang et al., 2005)

(McCullough et al., 1987; Liang et al., 2005)

(Carter et al., 2000; Laird et al., 2001)

(Ok et al., 2006)

VIPR ¼ 1.3, EP Kr ¼ 10, Ki ¼ 0.14 mM

VIPR IC50 ¼ 4

Binding Ki ¼ 49 nM, IC50i ¼ 77 nM, IC50r ¼ 18 mM

EP IC50 ¼ 0.64, 0.36 (Nav1.7)

Nav1.7 block, others unknown

Nav1.7 block, others unknown

Nonselective

Unknown

71

Unknown

230

Unknown

(Continued)

Formalin

Formalin (phase 2); s.c. only

CFA

CFA

372

A-803467

4030W92

M58373

12

Name

CN

Cl

TABLE 14.2

Cl

N

Cl

O

H2N

O

HCl

N

O

H

N

N

O

NH

N

OH

S

Structure

(Continued )

F

N

O

NH2

O

O

Cl

(Jarvis et al., 2007)

(Clayton et al., 1998; Collins et al., 1998; Trezise et al., 1998)

(Akada et al., 2006)

(Liang et al., 2005)

References

EP IC50 10 nM

EP IC50 ¼ 22 at
60 mV (TTX-R) and 5 at
70 mV (TTX-S)

Binding Ki ¼ 0.7

VIPR ¼ 1.0 EP Kr ¼ 6.7, Ki ¼ 0.16 mM

Potency (mM)

Nav1.8  Nav1.5, Nav1.2, Nav1.7

Nonselective

Unknown

Nav1.7 block, others unknown

Selectivity (Most to Least Potent)

Formalin

Pain Models (In Vivo)

10

Yes

CFA, CCI, capsaicin, SNL, SNI. inactive in formalin

Formalin (phase 2), carrageenan, CFA, CCI

Unknown Formalin, CCI

42

UseDependent Ratio

373

#47

PPPA

Cl

N

O

NH

O O

F

O

O

NH

F3CO

N

N

O

NH2

(Hoyt et al., 2007)

(Ilyin et al., 2006)

EP IC50 ¼ 35 nM Ki ¼ 40 nM, Kr >3 mM

EP Ki ¼ 0.041, Kr ¼ 22 mM

Nav1.7 block, others unknown

Nonselective

>75

537

SNL tactile allodynia (10 mg/kg p.o.)

Partial sciatic nerve ligation, CFA, postincisional (mechanical endpoints)

374

14.2.3

SODIUM CHANNEL BLOCKERS FOR THE TREATMENT OF CHRONIC PAIN

Local Anesthetics

Lidocaine (lignocaine, xylocaine) was first synthesized in 1943 and is still the most widely used local anesthetic. Although lidocaine is a relatively weak blocker of Naþ channels (IC50 generally 50 mM), it does block in a state-dependent manner (approximately fivefold more potent (Smith et al., 2006)), which allows neuronal stabilization without blocking conduction (see Table 14.1). Lidocaine also preferentially blocks TTX-R currents (Roy and Narahashi, 1992; Chevrier et al., 2004; Weiser, 2006), which are predominantly expressed in primary afferent neurons, a feature that may contribute to lidocaine’s discrimination between sensory and motor effects. Due to its poor oral bioavailability, lidocaine must be injected locally or infused as an intravenous or epidural solution. In rat pain models, systemic lidocaine has been shown to be effective at preventing pain behaviors following formalin injection and also in reversing thermal hyperalgesia associated with sciatic nerve ligation without affecting processing on the uninjured side (Abram and Yaksh, 1994). Lidocaine has been extensively studied in human clinical trials (for reviews see Kalso et al., 1998; Tremont-Lukats et al., 2005) and has been shown to be broadly effective against pain associated with amputation (Wu et al., 2002), spinal cord injury (Finnerup et al., 2005), and postherpetic neuralgia (intravenous) (Rowbotham et al., 1991) among others. Lidocaine (Lidoderm) administered topically as a patch is only one of the five drugs currently approved by the Food and Drug Administration (FDA) for the treatment of chronic pain. Although the lidocaine patch is approved for postherpetic neuralgia, pilot studies suggest that it may also be effective for alleviating the pain and potentially other symptoms associated with osteoarthritis, lower back pain, and diabetic neuropathy (Argoff et al., 2004; Burch et al., 2004; Gimbel et al., 2005). The nonselective nature of the Naþ channel block by lidocaine, however, can lead to adverse events and doselimiting side effects (Challapalli et al., 2005; Tremont-Lukats et al., 2005). Mexiletine was developed as an orally available analogue of lidocaine. It is also a state-dependent (three-fourfold) (De Luca et al., 2000) and relatively weak blocker of Naþ channels with in vitro potencies in the mM range, although it is slightly more potent than lidocaine. Mexiletine is currently approved by the FDA only for the treatment of life-threatening ventricular arrhythmia and not for the treatment of pain. Mexiletine is effective in reducing mechanical allodynia induced by peripheral nerve injury in animal models of neuropathic pain (Erichsen and Blackburn-Munro, 2002; Lindia et al., 2005) and in reducing flinching behavior in both phase 1 and phase 2 of the formalin test (Blackburn-Munro et al., 2002). There are human studies examining the efficacy of oral mexiletine in chronic pain compared to lidocaine (usually i.v.), and the results are mixed. Mexiletine has been shown to be effective in pain associated with peripheral nerve damage (Chabal et al., 1992) and diabetic neuropathy (Dejgard et al., 1988), but ineffective against central pain associated with spinal cord injury (ChiouTan et al., 1996), HIV-related painful peripheral neuropathy (Kemper et al., 1998), and others (Ando et al., 2000; Wallace et al., 2000). Side effects and adverse events associated with systemic mexiletine in these studies were generally mild and included nausea, gastrointestinal disturbances, dizziness, and tremor.

PHARMACOLOGY

14.2.4

375

Antiepileptics

Many antiepileptic drugs such as carbamazepine and lamotrigine have Naþ channel blocking activity, which contributes to their antiepileptic activity (see Table 14.1). Generally these are weak but use-dependent blockers of Naþ channels, although other activities of these compounds, such as block of voltage-gated Ca2þ channels (Stefani et al., 1996), may contribute to their in vivo profile. Carbmazepine and lamotrigine block both TTX-S and TTX-R Naþ channels in DRG neurons with low affinity but show a strong preference for the inactivated state (inactivated state affinities of 28 mM and 111 mM, respectively, for carbamazepine) (Ilyin et al., 2006). In the rat formalin model, both carbamazepine and lamotrigine reduce phase 2 flinching behavior (but not phase 1) at doses that do not affect motor coordination (Blackburn-Munro et al., 2002). Results for carbamazepine and lamotrigine in animal models of neuropathic pain are mixed and appear to be model dependent and sedative, or ataxic side effects can be seen at or near the efficacious doses (Ilyin et al., 2006). Clinically, carbamazepine is primarily used as an antiepileptic, although it is also approved by the Food and Drug Administration for the treatment of pain, where it is the first line treatment for trigeminal neuralgia. Evidence from clinical trials also suggests the utility of carbamazepine in the treatment of painful diabetic neuropathy (Gomez-Perez et al., 1996; Vinik, 2005). Overall the clinical data on lamotrigine is inconclusive and lamotrigine is only considered a fourth line treatment for neuropathic pain (Moulin et al., 2007). Zonisamide is another antiepileptic drug that has proven useful in treating chronic pain. A number of activities have been attributed to zonisamide including Naþ channel blockade, Cav channel blockade, potassium channel potentiation, nitric oxide synthase inhibition, and nitric oxide free radical scavenger activity. Many of these activities may contribute to antinociceptive action, making it difficult to assign a clear mechanism of action to zonisamide in in vivo studies. Given this caveat, systemically administered zonisamide has been shown to alleviate acute thermal pain in mice (Sakaue et al., 2004) and is effective at relieving thermal hyperalgesia and tactile allodynia in nerve injury models of chronic pain (Hord et al., 2003; Tanabe et al., 2005). There are a number of clinical case studies and open-label reports suggesting that zonisamide is effective in alleviating chronic pain in humans; however, data from few randomized controlled trials have been published (Guay, 2003). Data from one randomized, placebo-controlled study examined 42 patients with painful diabetic neuropathy. Pain scores for the zonisamide-treated group decreased more than the placebo group, although these differences did not reach statistical significance (Atli and Dogra, 2005). The most unusual member of this category may well be lacosamide (harkoseride or ADD 234037), a functionalized amino acid that was specifically designed as an anticonvulsant. Unlike the other Naþ channel blocking compounds, lacosamide selectively enhances slow inactivation of Naþ channels (and binds collapsin-response mediator protein 2 or CRMP-2) without affecting fast inactivation (Errington et al., 2006; Beyreuther et al., 2007b). Some other anticonvulsant compounds such as zonisamide are reported to enhance fast and slow inactivation (Schauf, 1987). In animal models of pain, lacosamide is generally without effect in acute pain models, but

376

SODIUM CHANNEL BLOCKERS FOR THE TREATMENT OF CHRONIC PAIN

inhibits chronic inflammatory and neuropathic pain. Activity has been reported in diverse thermal and mechanical endpoints in the inflammatory carrageenan and CFA models and in the models of streptocotozin-induced diabetic neuropathic pain, arthritic pain, and spinal cord and infraorbital injury-induced pain (Beyreuther et al., 2006, 2007a, 2007c; Hao et al., 2006; Stohr et al., 2006). Thus, lacosamide has broad antinociceptive activities in chronic pain models. Clinical experience has been encouraging as well; patients with painful diabetic neuropathy have reported strong and sustained pain reduction in several multicenter, randomized, double-blind, placebo-controlled trials (McCleane et al., 2003; Shaibani et al., 2006; Wymer et al., 2006; Rauck et al., 2007).

14.3 NEW SMALL-MOLECULE PHARMACOLOGY Many small-molecule Nav blockers have been identified as binding to the “local anesthetic” binding site of Naþ channels. It is apparent that this site is relatively promiscuous with respect to the binding of small drug-like molecules and is partially conserved across all the Nav subtypes. Many investigators have used the available pharmacological agents such as batracotoxin (BTX), veratridine, and other tools which open or prevent inactivation of Nav channels, enabling assays for Nav channel inhibitors on multiwell higher throughput platforms where membrane potential cannot be tightly controlled. Compounds that bind to the local anesthetic site, as it turns out, also allosterically inhibit BTX binding to its site, thus providing the basis for a binding assay for the discovery of blocking compounds (Postma and Catterall, 1984; Sheldon et al., 1994). This fact has likely contributed to the large variety of modulators discovered in past years, as classical ligand binding assays based on the displacement of radiolabeled BTX have proven useful for identifying “hits.” One limitation of this method, however, is that most studies have used brain or heart tissue/channels; thus, the compounds identified are not selective for channels specifically involved in pain states. Another limitation is that such a method discovers modulators that act in a manner similar to previous compounds, in this case those that inhibit BTX binding and are therefore likely binding to the local anesthetic site. Nevertheless, the newer pharmacological agents have generally evolved from these nonselective starting points, and other plate-based screens have been performed utilizing the pain-related Naþ channels Nav1.7 and Nav1.8, minimizing the first of these limitations. The activity of these new agents on Nav channels and in subsequent pain models has built a compelling picture for the effectiveness of Naþ channel blockers in treating pain in recent years and is detailed in the following sections and in Table 14.2. 14.3.1

Ralfinamide (NW-1029)

A series of largely nonselective Nav blockers that inhibit binding to the batrachotoxinin site (neurotoxin 2) in native DRG cells is exemplified by NW-1029 (ralfinamide). NW-1029 blocks TTX-S and TTX-R channels in DRG neurons in a use-dependent

NEW SMALL-MOLECULE PHARMACOLOGY

377

fashion, although its potency is unimpressive. At physiological membrane potentials of
70 mV, the IC50 ¼ 55 mM for TTX-R and 22 mM for TTX-S channels, but 50 mM is able to shift the V1/2 for steady-state inactivation by 11 and 15 mV for TTX-R and TTX-S channels, respectively (Stummann et al., 2005). Ralfinamide strongly blocked the phase 2 (neuropathic-like) pain in the formalin model (10–40 mg/kg p.o.) and produced some decrease in phase 1 (acute inflammatory) pain at 40 mg/kg p.o. In the inflammatory CFA-induced mechanical allodynia model, activity was observed with an ED50 of 0.5 mg/kg i.p. or p.o. In the chronic constriction injury (CCI) rat model of chronic inflammatory pain, mechanical allodynia was blocked with ED50s of 0.9 and 0.7 mg/kg i.p. and p.o., respectively (Veneroni et al., 2003). No effect was observed in acute pain models of hot-plate and tail-flick tests in the rat, whereas in the rotarod test of motor coordination, TD50s of 245–470 mg/kg p.o. predict a wide therapeutic index for this molecule. Suppression of seizures was also demonstrated for NW-1029 and its related molecules in a variety of anticonvulsant models (Pevarello et al., 1998), thus supporting the link between hyperexcitability in chronic pain and epilepsy models. In rats, ralfinamide was effective in reducing spontaneous neuropathic pain behavior, via its suppression of self-amputation (autotomy). Compared with vehicle-treated rats, ralfinamide significantly delayed the autotomy onset day in a dose-dependent manner over a 42-day study. Autotomy levels in rats treated with a 120 mg/kg/day dose remained significantly suppressed until the end of the experiment (postoperative day 63). In a parallel electrophysiological study, it was shown that ralfinamide blocked ectopic neuroma discharges generated in sensory fibers without affecting normal nerve conduction (Seltzer et al., 2006). Clinical experience with this compound has progressed as far as phase 2, where ralfinamide produced a significant improvement in pain scores as measured by the visual analogue scale of pain intensity (VASPI; 26% improvement versus baseline). The trial enrolled 18 patients who were administered ascending oral doses of ralfinamide (80, 160, 320 mg/day) for 4 weeks. Pain relief was experienced in >70% of patients in at least one category. Pin-prick allodynia improved in 67% of patients (12/18) and in seven patients (39%) evoked pain was reduced by 50%. Improvements were also observed in several other global pain scales and measures (Gustorff and Palmas, 2005). Avery structurally similar compound safinamide is also a Naþ channel blocker, but it also possesses monoamine oxidase-B and Cav inhibitory activity. This compound has been investigated as an antiepileptic and Parkinson’s disease treatment, but may be effective for pain as well (Fariello et al., 1998; Pevarello et al., 1998; Salvati et al., 1999). 14.3.2

BPBTS, CDA54

High-throughput screening for blockers of the peripherally expressed and pain implicated Nav1.7 have been performed using a fluorescent, functional assay designed to detect the voltage change in a Nav1.7-expressing cell line induced upon addition of Naþ channel openers. Using FRET and VIPR technologies yielded a compound termed “BPBTS” (Felix et al., 2004; Ok et al., 2006). BPBTS is

378

SODIUM CHANNEL BLOCKERS FOR THE TREATMENT OF CHRONIC PAIN

significantly more potent than other Naþ channel blockers currently used to treat neuropathic pain, although like these other agents, its block is largely nonselective across the different Nav subtypes (Priest et al., 2004). Block by BPBTS was found to be voltage and use dependent, with binding preferentially to open and inactivated states of the channel, which causes a dose-dependent hyperpolarizing shift in the steady-state availability curves for all the sodium channel subtypes tested. Although BPBTS has poor pharmacokinetic characteristics, it was still able to reduce the early and late phase pain behavior in the formalin model (Priest et al., 2004; Shao et al., 2005). Medicinal chemistry efforts to optimize BPBTS have led to CDA54, a more metabolically stable analogue that also blocks Nav1.8/1.7, but in an even greater usedependent manner (Shao et al., 2005). The potency is similar to BPBTS, but block of the pain-related Nav1.8 is enhanced (although block of cardiac-expressed Nav1.5 is still prominent) (Brochu et al., 2006). To characterize its activity in a chronic nerve injury model, the CCI model was examined. In this model, the induced injury leads to spontaneous action potential firing in the sciatic nerve, and block of these ectopic discharges is thought to at least partially underlie relief from neuropathic pain. CDA54 was able to block these spontaneous action potentials at 10-fold lower concentrations than those associated with conduction block in A or C fibers, thus dissociating potential analgesic and antihyperalgesic effects. Pain relief is also produced in CCI as well as in the SNL neuropathic model, at 10 mg/kg p.o. in both models. Unlike most other compounds of this type, when dosed in vivo, the distribution of CDA54 is largely restricted to peripheral tissues, yet it is still active in the formalin model, reducing pain behaviors to a similar level as that observed with mexiletine. Consistent with the spinal nerve data, no effect was found on acute nociception, motor coordination, or cardiac electrophysiology, except at higher concentrations (Shao et al., 2005; Brochu et al., 2006). Recently, further improvements in BPBTS and CDA54 have been made, especially targeting the metabolic instability of the two former compounds upon incubation with human liver microsomes. The more metabolically stable compounds 13 (racemate) and 13B (utamer, i.e., most active) were described to have similar potency as CDA54, better selectivity versus the hERG cardiac potassium channel, and thus were tested in vivo. The acute inflammatory CFA pain model was examined, where CDA54, 13, and 13B were all shown to have similar antiallodynic activity at 3 mg/kg p.o., thus indicating the therapeutic potential for these blockers (Ok et al., 2006). 14.3.3

Crobenetine (BIII 890 CL)

Discovered by investigators at Boehringer Ingelheim and the University of Washington, the compound crobenetine (BIII 890 CL) reduced binding of [3H]BTX A-20abenzoate at the neurotoxin receptor site 2 of the mixed Naþ channels expressed in rat brain synaptosomes (IC50 ¼ 49 nM) and TTX-R Nav channels in DRG (Carter et al., 2000; Dekker et al., 2005). A strong preference for the inactivated state (ratio of 230 in the IC50s for the resting and inactivated states) and use dependence has been described

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for its action. A further exploration of the binding site has been performed using sitedirected mutagenesis of cloned channels, where an alteration in compound binding occurred when channels were mutated in the local anesthetic site, indicating that binding occurs to this site (Carter et al., 2000). In rat cortical and striatal brain slices, 0.3 mM crobenetine inhibited veratridine-induced glutamate release, whereas in serum-free cultures of cortical neurons, it inhibited veratridine-induced glutamate release and neurotoxicity/cytotoxicity (Hamasaki et al., 1996; Carter et al., 2000). Crobenetine is active at blocking inflammatory pain in the CFA model in rats, with mechanical hyperalgesia and joint stiffness being significantly reduced after 5 days of treatment, with no effect on edema (Laird et al., 2001). These results were similar to those observed with mexiletine. Neuroprotective effects have also been reported at doses as low as 3 mg/kg s.c. in transient and permanent models of ischemic stroke in rats, whereas no effects were observed with 30 mg/kg s.c. in the rotarod test of motor coordination (Carter et al., 2000). 14.3.4

BA 42730A

A blocker of cardiac Naþ current described in an abstract (McCullough et al., 1987) as BA 42730A was shown recently to also block Nav1.7 and have activity in phase 2 of formalin-induced pain (Liang et al., 2005). This compound, however, was not selective for Nav channels, as it blocked other channels including hERG. Improved analogues (compounds 11 and 12) (Liang et al., 2005) were thus produced, which contained little hERG activity while retaining Nav1.7 block. These analogues produced significant activity in the formalin model of pain. Their potency on other Nav channels is unknown, but it is unlikely that these compounds are selective for Nav1.7. 14.3.5

M58373

In the veratridine-induced cytotoxicity assay (Hamasaki et al., 1996), M58373 was identified as a potent inhibitor of cell death (Akada et al., 2006). Determined to have an affinity (Ki) of 0.7 mM in the [3H]BTX binding assay subsequent experiments determined that M58373 blocked veratridine-induced release of substance-P from DRG neurons. In the formalin test, oral M58373 (0.3–10 mg/kg) reduced nociceptive behaviors in phase 2, while in the CCI chronic pain model, oral M58373 (1–10 mg/kg) attenuated mechanical allodynia and thermal hyperalgesia without affecting normal responses in the uninjured paw (Akada et al., 2006). 14.3.6

4030W92

Starting with the antiepileptic drug lamotrigine, the novel compound 4030W92 has been described as a voltage- and use-dependent Naþ channel inhibitor (Trezise et al., 1998). Both TTX-R and TTX-S channels are blocked in rat DRG small-diameter neurons in vitro, showing a preferential affinity (five–seven fold) for the slow inactivated state of the channel. Although a weak blocker with only micromolar activity, potential apparently exists for the treatment of chronic inflammatory (Clayton et al., 1998) and neuropathic

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(Collins et al., 1998) pain, although much of the data has been published only in abstract form. This compound was examined in a clinical trial of 41 subjects with chronic neuropathic pain with prominent allodynia (Wallace et al., 2002). 4030W92 (25 mg/day) reduced allodynia on day 1 but not on days 7 or 14 of a 2 week trial. 14.3.7

A-803467

A new compound A-803467, discovered and characterized collaboratively by Abbott and ICAgen, has been described recently (Jarvis et al., 2007). It is the most selective compound described for DRG-associated Nav1.8 channels, use-dependently blocking with an IC50 of 10 nM while only blocking other Nav channels (Nav1.2, 1.5, 1.7) with IC50s of 3–10 mM. This activity profile translates into a dose-dependent reduction of mechanical allodynia in both neuropathic and inflammatory models (SNL, SNI, capsaicin-induced secondary mechanical allodynia and thermal hyperalgesia following CFA). A-803467 was inactive in the formalin, acute and postoperative pain models, and also showed no deficits in spontaneous exploratory behaviors or motor coordination. Thus, specific Nav1.8 blockade appears to result in broad activity across neuropathic and inflammatory models, while showing little or no effect on motor coordination or exploratory activity and no effects on other behavioral measures of peripheral nerve activity, including acute pain. Thus, pharmacological blockade of Nav1.8 appears to be an efficacious and tolerated method for the relief of chronic pain, at least in these animal models. 14.3.8

PPPA

V102862 was originally described as an anticonvulsant (Dimmock et al., 1993, 1996) and subsequently discovered to be a broad-spectrum, state-dependent blocker of voltage-gated Naþ channels (Ilyin et al., 2005). In an effort to obtain control over pain states and a better therapeutic index, broad-spectrum Naþ channel blockers with higher potency, faster onset kinetics, and greater state dependence than existing drugs were the goal of an optimization effort around V102862 (Shao et al., 2004; Yang et al., 2004). The resultant compound, PPPA, met these criteria, displaying greater than 1000-fold better potency and binding kinetics and 10-fold more state dependence than the comparators carbamazepine and lamotrigine (Ilyin et al., 2006). This activity translated into activity in the partial sciatic nerve ligation, CFA, and postincisional pain models using mechanical endpoints, although efficacy was similar to carbamazepine and lamotrigine. With a therapeutic index >10 when measured by ataxia in the rotarod model, PPPA demonstrated better tolerability than the comparator compounds, thus indicating that this approach may prove effective for identifying new Nav channel blockers for pain. 14.3.9

Merck Compound 47

Last, a potent, state-dependent Nav1.7 blocker termed #47 (Hoyt et al., 2007) has been described in a medicinal chemistry-focused paper. Neuronal firing in vivo

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was inhibited in the rat peripheral axotomy model with 2 mg/kg i.v., while oral activity (10 mg/kg) was observed in the SNL tactile allodynia model. Further optimization of pharmacokinetic and pharmacodynamic parameters reportedly continues.

14.4 ANTIDEPRESSANTS Tricyclic antidepressants (TCAs) such as amitriptyline have long been used to treat chronic pain conditions (see Table 14.1), either alone or as adjuvant treatment for opioid therapy. Administration of antidepressants has been shown to reduce pain in patients with diabetic neuropathy (Max et al., 1987, 1992a), post herpetic neuralgia (Watson et al., 1982), and other chronic pain conditions (Magni, 1991; Onghena and Van Houdenhove, 1992; Finnerup et al., 2005). Drugs in this class are usually thought to alleviate pain by increasing levels of norepinephrine and serotonin, thereby stimulating descending inhibitory pain pathways. However, many of the compounds in this class also exhibit Nav, Cav, and ionotropic glutamate receptor-blocking activities, which may contribute to their antinociceptive profile. Amitriptyline is the most widely prescribed TCA for the treatment of chronic pain. Many reports describe the inhibitory action of amitriptyline on Naþ channels in vitro and it is generally described as a nonselective, use-dependent blocker with moderate potency (IC50 from 5–80 mM). In isolated rat DRG neurons, amitriptyline inhibited both the fast (TTX-S) and slow (TTX-R) sodium currents with similar potency (IC50 20 mM) (Pancrazio et al., 1998). A recent paper (Dick et al., 2007) examined the blockade of the peripheral sodium channel Nav1.7 by amitriptyline in detail, and reports that the affinity for the inactivated state of the channel is 0.24 mM, a value that is less than the clinical plasma concentration for amitriptyline. Efficacy in the in vivo neuropathic SNL model has been demonstrated (Mochizucki, 2004), which has translated into clinical efficacy in diabetic neuropathy (Joss, 1999) and postherpetic neuralgia (Watson et al., 1982; Watson and Evans, 1985). Imipramine, another widely prescribed TCA, has long been known to have Naþ channel blocking properties (Schauf et al., 1975). Imipramine is a use-dependent blocker that preferentially binds to and stabilizes the inactivated state of Naþ channels (Dick et al., 2007), although an additional open-channel blocking mechanism has been described (Yang and Kuo, 2002). Imipramine has been demonstrated to be effective in certain human experimental pain models (Bromm et al., 1986; Poulsen et al., 1995; Enggaard et al., 2001) and in patients with painful peripheral polyneuropathy, including diabetic neuropathy (Kvinesdal et al., 1984; Sindrup et al., 2003). Although numerous reports demonstrate the clinical efficacy of TCAs against chronic pain, the preclinical efficacy data for both amitriptyline and imipramine are mixed. Imipramine was shown to be ineffective at reducing tactile and cold allodynia in a number of models of neuropathic pain (Hama and Borsook, 2005) and the

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hot-plate model of acute thermal pain (Otsuka et al., 2001), although it was shown to be analgesic in an electrical shock model of acute pain (Abdel-Salam et al., 2003). There are numerous reports showing efficacy of imipramine and other TCAs in the rat formalin model. The effects of TCAs in the formalin model are largely reversed by noradrenergic and serotonergic antagonists, suggesting that Naþ channel blockade, however, is not the dominant mechanism of action for these compounds in this model (Otsuka et al., 2001; Yokogawa et al., 2002). The effectiveness of TCAs in pain conditions prompted the examination of other antidepressants in pain. Serotonin-norepinephrine reuptake inhibitors (SNRIs) are generally effective in chronic pain models (Iyengar et al., 2004; Mochizucki, 2004) and clinically in diabetic neuropathy (Max et al., 1992). In contrast, selective serotonin reuptake inhibitors (SSRIs) have been shown to be ineffective clinically in chronic pain conditions (Watson and Evans, 1985; Max et al., 1992b). Recently, however, all three types of antidepressant have been shown to block Naþ channels in a use-dependent manner (Huang et al., 2006). The therapeutic concentrations (for their effective use in depression) of the SSRIs, however, are lower than their affinity for the inactivated state of Nav1.7, whereas that of the TCAs and NRIs are greater than their affinity for Nav1.7 (Dick et al., 2007). Thus, the activity of antidepressants in pain conditions may be more related to their Naþ channel blocking activity than to their activity at inhibiting monoamine reuptake. This supports the role of Naþ channel blocking activity in the pain efficacy displayed by other antidepressants.

14.5 CONCLUSIONS Experience over the past half-century has demonstrated repeatedly the therapeutic value of small-molecule blockers of Naþ channels for many disorders of neuronal hyperexcitability. Recent years have seen a resurgence of interest in Nav channel blockers for chronic pain conditions, especially focused around Nav1.3 and the peripherally expressed Nav1.7–1.9 varieties. While many programs have focused on obtaining strongly use-dependent blockers as a means of targeting hyperexcitable cells selectively, recent successes have been notable for their ability to discover Nav subtype selective blockers, a feat previously thought unattainable. The recent recognition that commonly used antidepressants also contain Naþ channel blocking activity may point to another potential use for this class of molecules in future therapeutic settings. The “best” Nav subtype to target for pain conditions has implicated Nav1.3, Nav1.8/1.9, and Nav1.7, but it is likely that broader testing in animal models and in human pain conditions will be the ultimate factor in deciding which subtype(s) are best to target for a particular type or category of pain condition. Indeed, if successful, multiple types of Nav blockers with different combinations of selectivity and use dependence will be needed clinically to allow for choice by individual patients and their medical providers, similar to how epilepsy and depression are treated currently. If this goal is realized, the efforts of many scientists will be a success and bring immeasurable benefits to many patients in need.

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ABBREVIATIONS TTX-S TTX-R DRG Kr Ki VIPR EP IC50i IC50r MIA STZ ION CFA CCI SNI SNL TD50 Cav CRMP-2 FRET

(tetrodotoxin-sensitive) (tetrodotoxin-resistant) (dorsal root ganglia) (association rate constant, resting state) (association rate constant, inactivated state) (voltage ion probe reader) (electrophysiology) (IC50 for inactivated state) (IC50 for resting state) (monosodium iodoacetate) (streptocotozin) (infraorbital nerve) (complete Freund’s adjuvant) (chronic constriction injury) (spinal (spared) nerve injury) (spinal nerve ligation) (toxic dose for 50% of subjects) (voltage-dependent Ca2þ channel) (collapsin-response mediator protein 2) (fluorescence resonance energy transfer)

REFERENCES Abdel-Salam OM, Nofal SM, El-Shenawy SM, 2003. Evaluation of the anti-inflammatory and anti-nociceptive effects of different antidepressants in the rat. Pharmacol Res 48: 157–165. Abram SE, Yaksh TL, 1994. Systemic lidocaine blocks nerve injury-induced hyperalgesia and nociceptor-driven spinal sensitization in the rat. Anesthesiology 80: 383–391. (Discussion 325A) Akada Y, Ogawa S, Amano K-i, Fukudome Y, Yamasaki F, Itoh M, Yamamoto I, 2006. Potent analgesic effects of a putative sodium channel blocker M58373 on formalin-induced and neuropathic pain in rats. Eur J Pharmacol 536: 248–255. Ambrosio AF, Soares-Da-Silva P, Carvalho CM, Carvalho AP, 2002. Mechanisms of action of carbamazepine and its derivatives, oxcarbazepine, BIA 2-093, and BIA 2-024. Neurochem Res 27: 121–130. Ando K, Wallace MS, Braun J, Schulteis G, 2000. Effect of oral mexiletine on capsaicininduced allodynia and hyperalgesia: a double-blind, placebo-controlled, crossover study. Reg Anesth Pain Med 25: 468–474. Argoff CE, Galer BS, Jensen MP, Oleka N, Gammaitoni AR, 2004. Effectiveness of the lidocaine patch 5% on pain qualities in three chronic pain states: assessment with the Neuropathic Pain Scale. Med Res Opin 20 (Suppl 2): S21–28.

384

SODIUM CHANNEL BLOCKERS FOR THE TREATMENT OF CHRONIC PAIN

Atli A, Dogra S, 2005. Zonisamide in the treatment of painful diabetic neuropathy: a randomized, double-blind, placebo-controlled pilot study. Pain Med 6: 225–234. Backonja MM, 1994. Local anesthetics as adjuvant analgesics. J Pain Symptom Manage 9: 491– 499. Beyreuther B, Callizot N, Stohr T, 2006. Antinociceptive efficacy of lacosamide in a rat model for painful diabetic neuropathy. Eur J Pharmacol 539: 64–70. Beyreuther B, Callizot N, Stohr T, 2007a. Antinociceptive efficacy of lacosamide in the monosodium iodoacetate rat model for osteoarthritis pain. Arthritis Res Ther 9: R14. Beyreuther BK, Freitag J, Heers C, Krebsfanger N, Scharfenecker U, Stohr T, 2007b. Lacosamide: a review of preclinical properties. CNS Drug Rev 13: 21–42. Beyreuther BK, Callizot N, Brot MD, Feldman R, Bain SC, Stohr T, 2007c. Antinociceptive efficacy of lacosamide in rat models for tumor- and chemotherapy-induced cancer pain. Eur J Pharmacol 565: 98–104. Blackburn-Munro G, Ibsen N, Erichsen HK, 2002. A comparison of the anti-nociceptive effects of voltage-activated Naþ channel blockers in the formalin test. Eur J Pharmacol 445: 231–238. Brochu RM, Dick IE, Tarpley JW, McGowan E, Gunner D, Herrington J, Shao PP, Ok D, Li C, Parsons WH, Stump GL, Regan CP, Lynch JJ Jr, Lyons KA, McManus OB, Clark S, Ali Z, Kaczorowski GJ, Martin WJ, Priest BT, 2006. Block of peripheral nerve sodium channels selectively inhibits features of neuropathic pain in rats. Mol Pharmacol 69: 823–832. Bromm B, Meier W, Scharein E, 1986. Imipramine reduces experimental pain. Pain 25: 245– 257. Bulaj G, Zhang M-M, Green BR, Fiedler B, Layer RT, Wei S, Nielsen JS, Low SJ, Klein BD, Wagstaff JD, Chicoine L, Harty TP, Terlau H, Yoshikami D, Olivera BM, 2006. Synthetic muO-conotoxin MrVIB blocks TTX-resistant sodium channel NaV1.8 and has a longlasting analgesic activity. Biochemistry 45: 7404–7414. Burch F, Codding C, Patel N, Sheldon E, 2004. Lidocaine patch 5% improves pain, stiffness, and physical function in osteoarthritis pain patients. A prospective, multicenter, open-label effectiveness trial. Osteoarthritis Cartilage 12: 253–255. Carter AJ, Grauert M, Pschorn U, Bechtel WD, Bartmann-Lindholm C, Qu Y, Scheuer T, Catterall WA, Weiser T, 2000. Potent blockade of sodium channels and protection of brain tissue from ischemia by BIII 890 CL. Proc Natl Acad Sci USA 97: 4944–4949. Catterall WA, 1987. Common modes of drug action on NaC channels: local anesthetics, antiarrhythmics and anticonvulsants. Trends Pharmacol Sci 8: 57–65. Chabal C, Jacobson L, Mariano A, Chaney E, Britell CW, 1992. The use of oral mexiletine for the treatment of pain after peripheral nerve injury. Anesthesiology 76: 513–517. Challapalli V, Tremont-Lukats IW, McNicol ED, Lau J, Carr DB, 2005. Systemic administration of local anesthetic agents to relieve neuropathic pain. Cochrane Database Syst Rev: CD003345. Chevrier P, Vijayaragavan K, Chahine M, 2004. Differential modulation of Nav1.7 and Nav1.8 peripheral nerve sodium channels by the local anesthetic lidocaine. Br J Pharmacol 142: 576–584. Chiou-Tan FY, Tuel SM, Johnson JC, Priebe MM, Hirsh DD, Strayer JR, 1996. Effect of mexiletine on spinal cord injury dysesthetic pain. Am J Phys Med Rehabil 75: 84–87. Clayton NH, Collins SD, Sargent R, Brown T, Nobbs M, Bountra C, Trezise DJ, 1998. The effect of the novel sodium channel blocker 4030W92 in models of acute and chronic inflammatory pain in the rat. Br J Pharmacol 123: 79P.

REFERENCES

385

Collins SD, Clayton NM, Nobbs M, Bountra C, 1998. The effect of 4030W92, a novel sodium channel blocker, on the treatment of neuropathic pain in the rat. Br J Pharmacol 123: 16P. De Luca A, Natuzzi F, Desaphy JF, Loni G, Lentini G, Franchini C, Tortorella V, Camerino DC, 2000. Molecular determinants of mexiletine structure for potent and use-dependent block of skeletal muscle sodium channels. Mol Pharmacol 57: 268–277. Dejgard A, Petersen P, Kastrup J, 1988. Mexiletine for treatment of chronic painful diabetic neuropathy. Lancet 1: 9–11. Dekker LV, Daniels Z, Hick C, Elsegood K, Bowden S, Szestak T, Burley JR, Southan A, Cronk D, James IF, 2005. Analysis of human Nav1.8 expressed in SH-SY5Y neuroblastoma cells. Eur J Pharmacol 528: 52–58. Dick IE, Brochu RM, Purohit Y, Kaczorowski GJ, Martin WJ, Priest BT, 2007. Sodium channel blockade may contribute to the analgesic efficacy of antidepressants. Journal of Pain 8: 315– 324. Dimmock JR, Sidhu KK, Thayer RS, Mack P, Duffy MJ, Reid RS, Quail JW, Pugazhenthi U, Ong A, Bikker JA, Weaver DF, 1993. Anticonvulsant activities of some arylsemicarbazones displaying potent oral activity in the maximal electroshock screen in rats accompanied by high protection indices. J Med Chem 36: 2243–2252. Dimmock JR, Puthucode RN, Smith JM, Hetherington M, Quail JW, Pugazhenthi U, Lechler T, Stables JP, 1996. (Aryloxy)aryl semicarbazones and related compounds: a novel class of anticonvulsant agents possessing high activity in the maximal electroshock screen. J Med Chem 39: 3984–3997. Eisenberg E, Shifrin A, Krivoy N, Lamotrigine for neuropathic pain. 2005. Expert Rev Neurother 5: 729–735. Ekberg J, Jayamanne A, Vaughan CW, Aslan S, Thomas L, Mould J, Drinkwater R, Baker MD, Abrahamsen B, Wood JN, Adams DJ, Christie MJ, Lewis RJ, 2006. mO-conotoxin MrVIB selectively blocks Na1.8 sensory neuron specific sodium channels and chronic pain behavior without motor deficits. Proc Natl Acad Sci USA 103: 17030-17035. Enggaard TP, Poulsen L, Arendt-Nielsen L, Hansen SH, Bjornsdottir I, Gram LF, Sindrup SH, 2001. The analgesic effect of codeine as compared to imipramine in different human experimental pain models. Pain 92: 277–282. Erichsen HK, Blackburn-Munro G, 2002. Pharmacological characterisation of the spared nerve injury model of neuropathic pain. Pain 98: 151–161. Errington AC, Coyne L, Stohr T, Selve N, Lees G, 2006. Seeking a mechanism of action for the novel anticonvulsant lacosamide. Neuropharmacology 50: 1016–1029. Faravelli L, Maj R, Veneroni O, Fariello RG, Benatti L, Salvati P, 2000. NW-1029 is a novel Na1 channel blocker with analgesic activity in animal models. In: Society for Neuroscience 30th Annual Meeting, New Orleans, LA, p 1218. Fariello RG, McArthur RA, Bonsignori A, Cervini MA, Maj R, Marrari P, Pevarello P, Wolf HH, Woodhead JW, White HS, Varasi M, Salvati P, Post C, 1998. Preclinical evaluation of PNU151774E as a novel anticonvulsant. J Pharmacol Exp Ther 285: 397–403. Felix JP, Williams BS, Priest BT, Brochu RM, Dick IE, Warren VA, Yan L, Slaughter RS, Kaczorowski GJ, Smith MM, Garcia ML, 2004. Functional assay of voltage-gated sodium channels using membrane potential-sensitive dyes. Assay Drug Dev Technol 2: 260–268. Finnerup NB, Biering-Sorensen F, Johannesen IL, Terkelsen AJ, Juhl GI, Kristensen AD, Sindrup SH, Bach FW, Jensen TS, 2005. Intravenous lidocaine relieves spinal cord injury pain: a randomized controlled trial. Anesthesiology 102: 1023–1030.

386

SODIUM CHANNEL BLOCKERS FOR THE TREATMENT OF CHRONIC PAIN

Gimbel J, Linn R, Hale M, Nicholson B, 2005. Lidocaine patch treatment in patients with low back pain: results of an open-label, nonrandomized pilot study. Am J Ther 12: 311–319. Gomez-Perez FJ, Choza R, Rios JM, Reza A, Huerta E, Aguilar CA, Rull JA, 1996. Nortriptyline-fluphenazine vs. carbamazepine in the symptomatic treatment of diabetic neuropathy. Arch Med Res 27: 525–529. Guay DRP, 2003. Oxcarbazepine, topiramate, zonisamide, and levetiracetam: potential use in neuropathic pain. Am J Geriatr Pharmacother 1: 18–37. Gustorff B, Palmas M, 2005. A pilot study of ralfinamide in the treatment of patients with neuropathic pain. Neurology 64: 121. Hama AT, Borsook D, 2005. Behavioral and pharmacological characterization of a distal peripheral nerve injury in the rat. Pharmacol Biochem Behav 81: 170–181. Hamasaki K, Kogure K, Ohwada K, 1996. A biological method for the quantitative measurement of tetrodotoxin (TTX): tissue culture bioassay in combination with a water-soluble tetrazolium salt. Toxicon 34: 490–495. Hao J-X, Stohr T, Selve N, Wiesenfeld-Hallin Z, Xu X-J, 2006. Lacosamide, a new antiepileptic, alleviates neuropathic pain-like behaviors in rat models of spinal cord or trigeminal nerve injury. Eur J Pharmacol 553: 135–140. Hille B, 2001. Ionic Channels of Excitable Membranes. Sinauer Associates, Inc., Sunderland, MA. Hord AH, Denson DD, Chalfoun AG, Azevedo MI, 2003. The effect of systemic zonisamide (Zonegran) on thermal hyperalgesia and mechanical allodynia in rats with an experimental mononeuropathy. Anesth Analg 96: 1700–1706. Hoyt SB, London C, Gorin D, Wyvratt MJ, Fisher MH, Abbadie C, Felix JP, Garcia ML, Li X, Lyons KA, McGowan E, MacIntyre DE, Martin WJ, Priest BT, Ritter A, Smith MM, Warren VA, Williams BS, Kaczorowski GJ, Parsons WH, 2007. Discovery of a novel class of benzazepinone Nav1.7 blockers: potential treatments for neuropathic pain. Bioorg Med Chem Lett 17: 4630–4634. Huang C-J, Harootunian A, Maher MP, Quan C, Raj CD, McCormack K, Numann R, Negulescu PA, Gonzalez JE, 2006. Characterization of voltage-gated sodium-channel blockers by electrical stimulation and fluorescence detection of membrane potential. Nat Biotechnol 24: 439–446. Ilyin VI, Hodges DD, Whittemore ER, Carter RB, Cai SX, Woodward RM, 2005. V102862 (Co 102862): a potent, broad-spectrum state-dependent blocker of mammalian voltage-gated sodium channels. Br J Pharmacol 144: 801–812. Ilyin VI, Pomonis JD, Whiteside GT, Harrison JE, Pearson MS, Mark L, Turchin PI, Gottshall S, Carter RB, Nguyen P, Hogenkamp DJ, Olanrewaju S, Benjamin E, Woodward RM, 2006. Pharmacology of 2-[4-(4-chloro-2-fluorophenoxy)phenyl]-pyrimidine-4-carboxamide: a potent, broad-spectrum state-dependent sodium channel blocker for treating pain states. J Pharmacol Exp Ther 318: 1083–1093. Iyengar S, Webster AA, Hemrick-Luecke SK, Xu JY, Simmons RMA, 2004. Efficacy of duloxetine, a potent and balanced serotonin-norepinephrine reuptake inhibitor in persistent pain models in rats. J Pharmacol Exp Ther 311: 576–584. Jarvis B, Coukell AJ, 1998. Mexiletine. A review of its therapeutic use in painful diabetic neuropathy. Drugs 56: 691–707. Jarvis MF, Honore P, Shieh CC, Chapman M, Joshi S, Zhang X-F, Kort M, Carroll W, Marron B, Atkinson R, Thomas J, Liu D, Krambis M, Liu Y, McGaraughty S, Chu K, Roeloffs R,

REFERENCES

387

Zhong C, Mikusa J, Hernandez G, Gauvin D, Wade C, Zhu C, Pai M, Scanio M, Shi L, Drizin I, Gregg R, Matulenko M, Hakeem A, Gross M, Johnson M, Kennan Marsh K, Wagoner PK, Sullivan JP, Faltynek CR, Krafte DS, 2007. A-803467, a potent and selective Nav1.8 sodium channel blocker, attenuates neuropathic and inflammatory pain in the rat. Proc Natl Acad Sci USA 104: 8520–8525. Jensen TS, 2002. Anticonvulsants in neuropathic pain: rationale and clinical evidence. Eur J Pain 6 (Suppl A): 61–68. Joss JD, 1999. Tricyclic antidepressant use in diabetic neuropathy. Ann Pharmacother 33: 996– 1000. Kalso E, 2005. Sodium channel blockers in neuropathic pain. Curr Pharm Des 11: 3005– 3011. Kalso E, Tramer MR, McQuay HJ, Moore RA, 1998. Systemic local-anaesthetic-type drugs in chronic pain: a systematic review. Eur J Pain 2: 3–14. Keizer DW, West PJ, Lee EF, Yoshikami D, Olivera BM, Bulaj G, Norton RS, 2003. Structural basis for tetrodotoxin-resistant sodium channel binding by mu-conotoxin SmIIIA. J Biol Chem 278: 46805–46813. Kemper CA, Kent G, Burton S, Deresinski SC, 1998. Mexiletine for HIV-infected patients with painful peripheral neuropathy: a double-blind, placebo-controlled, crossover treatment trial. J Acquir Immune Defic Syndr Hum Retrovirol 19: 367–372. Kvinesdal B, Molin J, Froland A, Gram LF, 1984. Imipramine treatment of painful diabetic neuropathy. JAMA 251: 1727–1730. Laird JM, Carter AJ, Grauert M, Cervero F, 2001. Analgesic activity of a novel use-dependent sodium channel blocker, crobenetine, in mono-arthritic rats. Br J Pharmacol 134: 1742– 1748. Lei M, Jones SA, Liu J, Lancaster MK, Fung SSM, Dobrzynski H, Camelliti P, Maier SKG, Noble D, Boyett MR, 2004. Requirement of neuronal- and cardiac-type sodium channels for murine sinoatrial node pacemaking. J Physiol 559: 835–848. Liang J, Brochu RM, Cohen CJ, Dick IE, Felix JP, Fisher MH, Garcia ML, Kaczorowski GJ, Lyons KA, Meinke PT, Priest BT, Schmalhofer WA, Smith MM, Tarpley JW, Williams BS, Martin WJ, Parsons WH, 2005. Discovery of potent and use-dependent sodium channel blockers for treatment of chronic pain. Bioorg Med Chem Lett 15: 2943–2947. Lindia JA, Kohler MG, Martin WJ, Abbadie C, 2005. Relationship between sodium channel NaV1.3 expression and neuropathic pain behavior in rats. Pain 117: 145–153. Liu CN, Michaelis M, Amir R, Devor M, 2000a. Spinal nerve injury enhances subthreshold membrane potential oscillations in DRG neurons: relation to neuropathic pain. J Neurophysiol 84: 205–215. Liu CN, Wall PD, Ben-Dor E, Michaelis M, Amir R, Devor M, 2000b. Tactile allodynia in the absence of C-fiber activation: altered firing properties of DRG neurons following spinal nerve injury. Pain 85: 503–521. Lyu YS, Park SK, Chung K, Chung JM, 2000. Low dose of tetrodotoxin reduces neuropathic pain behaviors in an animal model. Brain Res 871: 98–103. Magni G, 1991. The use of antidepressants in the treatment of chronic pain. A review of the current evidence. Drugs 42: 730–748. Max MB, Culnane M, Schafer SC, Gracely RH, Walther DJ, Smoller B, Dubner R, 1987. Amitriptyline relieves diabetic neuropathy pain in patients with normal or depressed mood. Neurology 37: 589–596.

388

SODIUM CHANNEL BLOCKERS FOR THE TREATMENT OF CHRONIC PAIN

Max MB, Lynch SA, Muir J, Shoaf SE, Smoller B, Dubner R, 1992a. Effects of desipramine, amitriptyline, and fluoxetine on pain in diabetic neuropathy. N Engl J Med 326: 1250–1256 (see comment). Max MB, Lynch SA, Muir J, Shoaf SE, Smoller B, Dubner R, 1992b. Effects of desipramine, amitriptyline, and fluoxetine on pain in diabetic neuropathy. N Engl J Med 326: 1250–1256. McCleane G, Koch BD, Rauschkolb C, 2003. Does lacosamide have an analgesic effect in human neuropathic pain? An open label study. Neurosci Lett 352: 117–120. McCullough JR, Trube G, Conder ML, 1987. Blockade of cardiac sodium channels by low concentrations of BA 42730A. Biophys J 51: 260a. McIntosh JM, Hasson A, Spira ME, Gray WR, Li W, Marsh M, Hillyard DR, Olivera BM, 1995. A new family of conotoxins that blocks voltage-gated sodium channels. J Biol Chem 270: 16796–16802. Mochizucki D, 2004. Serotonin and noradrenaline reuptake inhibitors in animal models of pain. Hum Psychopharmacol 19 (Suppl 1): S15–S19. Moulin DE, Clark AJ, Gilron I, Ware MA, Watson CP, Sessle BJ, Coderre T, Morley-Forster PK, Stinson J, Boulanger A, Peng P, Finley GA, Taenzer P, Squire P, Dion D, Cholkan A, Gilani A, Gordon A, Henry J, Jovey R, Lynch M, Mailis-Gagnon A, Panju A, Rollman GB, Velly A, 2007. Pharmacological management of chronic neuropathic pain–consensus statement and guidelines from the Canadian Pain Society. Pain Res Manag 12: 13–21. Ogata N, Ohishi Y, 2002. Molecular diversity of structure and function of the voltage-gated Naþ channels. Jpn J Pharmacol 88: 365–377. Ok D, Li C, Abbadie C, Felix JP, Fisher MH, Garcia ML, Kaczorowski GJ, Lyons KA, Martin WJ, Priest BT, Smith MM, Williams BS, Wyvratt MJ, Parsons WH, 2006. Synthesis and SAR of 1,2-trans-(1-hydroxy-3-phenylprop-1-yl)cyclopentane carboxamide derivatives, a new class of sodium channel blockers. Bioorg Med Chem Lett 16: 1358–1361. Onghena P, Van Houdenhove B, 1992. Antidepressant-induced analgesia in chronic nonmalignant pain: a meta-analysis of 39 placebo-controlled studies. Pain 49: 205–219. Otsuka N, Kiuchi Y, Yokogawa F, Masuda Y, Oguchi K, Hosoyamada A, 2001. Antinociceptive efficacy of antidepressants: assessment of five antidepressants and four monoamine receptors in rats. J Anesth 15: 154–158. Pancrazio JJ, Kamatchi GL, Roscoe AK, Lynch C 3rd 1998. Inhibition of neuronal Naþ channels by antidepressant drugs. J Pharmacol Exp Ther 284: 208–214. Pevarello P, Bonsignori A, Dostert P, Heidempergher F, Pinciroli V, Colombo M, McArthur RA, Salvati P, Post C, Fariello RG, Varasi M, 1998. Synthesis and anticonvulsant activity of a new class of 2-[(arylalky)amino]alkanamide derivatives. J Med Chem 41: 579–590. Postma SW, Catterall WA, 1984. Inhibition of binding of [3H]batrachotoxinin A 20-alphabenzoate to sodium channels by local anesthetics. Mol Pharmacol 25: 219–227. Poulsen L, Arendt-Nielsen L, Brosen K, Nielsen KK, Gram LF, Sindrup SH, 1995. The hypoalgesic effect of imipramine in different human experimental pain models. Pain 60: 287–293. Priest BT, Garcia ML, Middleton RE, Brochu RM, Clark S, Dai G, Dick IE, Felix JP, Liu CJ, Reiseter BS, Schmalhofer WA, Shao PP, Tang YS, Chou MZ, Kohler MG, Smith MM, Warren VA, Williams BS, Cohen CJ, Martin WJ, Meinke PT, Parsons WH, Wafford KA, Kaczorowski GJ, 2004. A disubstituted succinamide is a potent sodium channel blocker with efficacy in a rat pain model. Biochemistry 43: 9866–9876.

REFERENCES

389

Ragsdale DS, McPhee JC, Scheuer T, Catterall WA, 1996. Proc Natl Acad Sci USA 93: 92. Rauck RL, Shaibani A, Biton V, Simpson J, Koch B, 2007. Lacosamide in painful diabetic peripheral neuropathy: a phase 2 double-blind placebo-controlled study. Clin J Pain 23: 150–158. Rowbotham MC, Reisner-Keller LA, Fields HL, 1991. Both intravenous lidocaine and morphine reduce the pain of postherpetic neuralgia. Neurology 41: 1024–1028. Roy ML, Narahashi T, 1992. Differential properties of tetrodotoxin-sensitive and tetrodotoxinresistant sodium channels in rat dorsal root ganglion neurons. J Neurosci 12: 2104–2111. Sakaue A, Honda M, Tanabe M, Ono H, 2004. Antinociceptive effects of sodium channelblocking agents on acute pain in mice. J Pharmacol Sci 95: 181–188. Salvati P, Maj R, Caccia C, Cervini MA, Fornaretto MG, Lamberti E, Pevarello P, Skeen GA, White HS, Wolf HH, Faravelli L, Mazzanti M, Mancinelli E, Varasi M, Fariello RG, 1999. Biochemical and electrophysiological studies on the mechanism of action of PNU-151774E, a novel antiepileptic compound. J Pharmacol Exp Ther 288: 1151– 1159. Schauf CL, 1987. Zonisamide enhances slow sodium inactivation in Myxicola. Brain Res 413: 185–188. Schauf CL, Davis FA, Kesler RL, 1975. Actions of the antidepressant drug imipramine on the voltage-clamped Myxicola giant axon. J Pharmacol Exp Ther 193: 669–675. Seltzer Z, Zhang S, Blech-Hermoni Y, Faravelli L, 2006. Ralfinamide suppresses spontaneous neuropathic pain behavior in the rat by differentially blocking ectopic ongoing neuroma discharges without affecting normal nerve conduction. Annual Meeting of the Society for Neuroscience, Atlanta, GA, 49.1/I12. Shaibani A, Kenney P, Simpson J, Bongardt S, 2006. Lacosamide in subjects with painful distal diabetic neuropathy: results of a multi-center, open-label, follow-on trial. J Pain 7: 43. Shao B, Victory S, Ilyin VI, Goehring RR, Sun Q, Hogenkamp D, Hodges DD, Islam K, Sha D, Zhang C, Nguyen P, Robledo S, Sakellaropoulos G, Carter RB, 2004. Phenoxyphenyl pyridines as novel state-dependent, high-potency sodium channel inhibitors. J Med Chem 47: 4277–4285. Shao PP, Ok D, Fisher MH, Garcia ML, Kaczorowski GJ, Li C, Lyons KA, Martin WJ, Meinke PT, Priest BT, Smith MM, Wyvratt MJ, Ye F, Parsons WH, 2005. Novel cyclopentane dicarboxamide sodium channel blockers as a potential treatment for chronic pain. Bioorg Med Chem Lett 15: 1901–1907. Sheldon RS, Duff HJ, Thakore E, Hill RJ, 1994. Class I antiarrhythmic drugs: allosteric inhibitors of [3H] batrachotoxinin binding to rat cardiac sodium channels. J Pharmacol Exp Ther 268: 187–194. Sindrup SH, Bach FW, Madsen C, Gram LF, Jensen TS, 2003. Venlafaxine versus imipramine in painful polyneuropathy: a randomized, controlled trial. Neurology 60: 1284–1289. Smith JA, Amagasu SM, Hembrador J, Axt S, Chang R, Church T, Gee C, Jacobsen JR, Jenkins T, Kaufman E, Mai N, Vickery RG, 2006. Evidence for a multivalent interaction of symmetrical, N-linked, lidocaine dimers with voltage-gated Naþ channels. Mol Pharmacol 69: 921–931. Sobieszek G, Borowicz KK, Bimber-Trojnar Z, Malek R, Piskorska B, Czuczwar SJ, 2003. Zonisamide: a new antiepileptic drug. Pol J Pharmacol 55: 683–689.

390

SODIUM CHANNEL BLOCKERS FOR THE TREATMENT OF CHRONIC PAIN

Stefani A, Spadoni F, Siniscalchi A, Bernardi G, 1996. Lamotrigine inhibits Ca2þ currents in cortical neurons: functional implications. Eur J Pharmacol 307: 113–116. Stohr T, Krause E, Selve N, 2006. Lacosamide displays potent antinociceptive effects in animal models for inflammatory pain. Eur J Pain 10: 241–249. Stummann TC, Salvati P, Fariello RG, Faravelli L, 2005. The anti-nociceptive agent ralfinamide inhibits tetrodotoxin-resistant and tetrodotoxin-sensitive Naþ currents in dorsal root ganglion neurons. Eur J Pharmacol 510: 197–208. Tanabe M, Sakaue A, Takasu K, Honda M, Ono H, 2005. Centrally mediated antihyperalgesic and antiallodynic effects of zonisamide following partial nerve injury in the mouse. Naunyn Schmiedebergs Arch Pharmacol 372: 107–114. Tanelian DL, Victory RA, 1995. Sodium channel-blocking agents: their use in neuropathic pain conditions. Pain Forum 4: 75–80. Tremont-Lukats IW, Challapalli V, McNicol ED, Lau J, Carr DB, 2005. Systemic administration of local anesthetics to relieve neuropathic pain: a systematic review and meta-analysis. Anesth Analg 101: 1738–1749. Trezise DJ, John VH, Xie XM, 1998. Voltage- and use-dependent inhibition of Naþ channels in rat sensory neurons by 4030W92, a new antihyperalgesic agent. Br J Pharmacol 124: 953–963. Veneroni O, Maj R, Calabresi M, Faravelli L, Fariello RG, Salvati P, 2003. Anti-allodynic effect of NW-1029, a novel Naþ channel blocker, in experimental animal models of inflammatory and neuropathic pain. Pain 102: 17–25. Vinik A, 2005. Clinical review: use of antiepileptic drugs in the treatment of chronic painful diabetic neuropathy. J Clin Endocrinol Metab 90: 4936–4945. Walia KS, Khan EA, Ko DH, Raza SS, Khan YN, 2004. Side effects of antiepileptics–a review. Pain Pract 4: 194–203. Wallace MS, Magnuson S, Ridgeway B, 2000. Efficacy of oral mexiletine for neuropathic pain with allodynia: a double-blind, placebo-controlled, crossover study. Reg Anesth Pain Med 25: 459–467. Wallace MS, Rowbotham M, Bennett GJ, Jensen TS, Pladna R, Quessy S, 2002. A multicenter, double-blind, randomized, placebo-controlled crossover evaluation of a short course of 4030W92 in patients with chronic neuropathic pain. J Pain 3: 227–233. Watson CP, Evans RJ, 1985. A comparative trial of amitriptyline and zimelidine in post-herpetic neuralgia. Pain 23: 387–394. Watson CP, Evans RJ, Reed K, Merskey H, Goldsmith L, Warsh J, 1982. Amitriptyline versus placebo in postherpetic neuralgia. Neurology 32: 671–673. Weiser T, 2006. Comparison of the effects of four Naþ channel analgesics on TTX-resistant Naþ currents in rat sensory neurons and recombinant Nav1.2 channels. Neurosci Lett 395: 179–184. West PJ, Bulaj G, Garrett JE, Olivera BM, Yoshikami D, 2002. Mu-conotoxin SmIIIA, a potent inhibitor of tetrodotoxin-resistant sodium channels in amphibian sympathetic and sensory neurons. Biochemistry 41: 15388–15393. Wiffen PJ, Rees J, 2007. Lamotrigine for acute and chronic pain. Cochrane Database Syst Rev CD006044. Wiffen PJ, McQuay HJ, Moore RA, 2005. Carbamazepine for acute and chronic pain. Cochrane Database Syst Rev CD005451.

REFERENCES

391

Wood JN, Boorman JP, Okuse K, Baker MD, 2004. Voltage-gated sodium channels and pain pathways. J Neurobiol 61: 55–71. Wu CL, Tella P, Staats PS, Vaslav R, Kazim DA, Wesselmann U, Raja SN, 2002. Analgesic effects of intravenous lidocaine and morphine on postamputation pain: a randomized double-blind, active placebo-controlled, crossover trial. Anesthesiology 96: 841–848. Wymer J, Garrison C, Simpson J, Koch B, 2006. A multicenter, randomized, double-blind, placebo-controlled trial to assess the efficacy and safety of lacosamide in subjects with painful distal diabetic neuropathy. Neurology 66: A202. Yang YC, Kuo CC, 2002. Inhibition of Naþ current by imipramine and related compounds: different binding kinetics as an inactivation stabilizer and as an open channel blocker. Mol Pharmacol 62: 1228–1237. Yang J, Gharagozloo P, Yao J, Ilyin VI, Carter RB, Nguyen P, Robledo S, Woodward RM, Hogenkamp DJ, 2004. 3-(4-Phenoxyphenyl)pyrazoles: a novel class of sodium channel blockers. J Med Chem 47: 1547–1552. Yokogawa F, Kiuchi Y, Ishikawa Y, Otsuka N, Masuda Y, Oguchi K, Hosoyamada A, 2002. An investigation of monoamine receptors involved in antinociceptive effects of antidepressants. Anesth Analg 95: 163–168 (Table of contents). Zorn S, Leipold E, Hansel A, Bulaj G, Olivera BM, Terlau H, Heinemann SH, 2006. The mOconotoxin MrVIA inhibits voltage-gated sodium channels by associating with domain-3. FEBS Lett 580: 1360–1364.

15 NEURONAL KV7 POTASSIUM CHANNELS AS EMERGING TARGETS FOR THE TREATMENT OF PAIN Steven I. Dworetzky and Valentin K. Gribkoff Knopp Neurosciences Inc., 2100 Wharton Street, Suite 615, Pittsburgh, PA 15203, USA

15.1 INTRODUCTION Potassium (Kþ) channels are the most diverse class of ion channels. Kþ currents are found in most cells and are associated with a wide range of functions, including the regulation of the electrical properties of excitable cells. The primary, pore-forming (a) subunits of these highly selective cation channels are divided into three primary structural classes based on the number of transmembrane (TM)-spanning regions and pore (P) regions: currently there are known to be 6TM/1P, 2TM/1P, and 4TM/2P Kþ channels. An additional 7TM representative that is a variant of the 6TM motif class has also been characterized (the BK channel, Slo1 or KCa1.1). Depending on the type of Kþ channel, the open probability (Popen) or activity of the particular voltagedependent Kþ channel can be regulated by transmembrane voltage (VM), protein phosphorylation and interaction with companion proteins as well as by a host of second messengers and endogenous ligands including ionic species such as calcium (Ca2þ), sodium (Naþ), and hydrogen (Hþ). The KV7 subfamily of voltage-gated Kþ channels consists of five homologous pore-forming a subunits, KV7.1–7.5, which have a structure typical of voltage-gated Kþ channels, with 6TM-spanning regions (S1–S6) flanked by intracellular N-terminal and C-terminal domains, a typical voltage-sensor domain located in S4 comprised of alternating positively charged

Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
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residues, and a single P region between S5 and S6 of each subunit (Fig. 15.1a). A subunit interaction and assembly complex is located in the cytoplasmic C-terminal domain (Howard et al., 2007; Wehling et al., 2007). The channels are formed as tetramers of the primary subunits, either as homotetramers or heterotetramers. The degree of homology between the members of the subfamily is shown in Fig. 15.1b.

FIGURE 15.1 (a) Illustration of the structure of a neuronal KV7 channel subunit. The KV7 protein has six transmembrane domains (S1–S6), a pore-forming P-loop, and a voltage sensor in the S4 domain indicated by the positively charged residues (þþþ). The channel is a tetramer of either homomultimers of the same subfamily member, such as KV7.2 channels comprising the substrate of neuronal IKS, or heteromultimeric mixtures of subfamily members, as is the case with M-channels that are heteromultimers of KV7.2/7.3 or KV7.5/7.3. The four alpha subunits (a1–a4) are indicated to illustrate the tetramerization of the channel complex, and the subunit interaction domains are also shown. Also illustrated is the relationship between neuronal KV7 channels and some receptor-mediated mechanisms of channel regulation. The (.) symbol in S5 marks the conserved amino acid between KV7.2–7.5 that is one known site of small-molecule interaction. (b) The table presents pair wise distances, shown as percent identity for purposes of illustration only, determined from the alignment of the human KV7.1–KV7.5 protein sequences. Factors that might affect these values include the inclusion of splice variants and the type of programs used to generate the alignment and the calculation of the sequence similarities.

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At least one member of the subfamily of KV7 channels is known to be associated with modulatory b subunits, an association that significantly affects the biophysical characteristics of the resulting currents and defines their physiological functions. KV7.1 associates with the b subunit KCNE1 to form the slow, depolarization-activated cardiac IKs current (Barhanin et al., 1996; Sanguinetti et al., 1996). The association of KV7.1 mutations with cardiac long-QT syndrome in humans resulted in its original designation as the KVLQT1 channel. In contrast, when the KV7.1 a subunit associates with the b subunit KCNE3 in intestinal crypt cells, the biophysical characteristics change to a constitutively open channel (Schroeder et al., 2000). The combination of KV7.1 with the b subunit KCNE2 in the gastric epithelium results in the channel responsible for coupling of Kþ to gastric acid secretion via colocalization of the channel with the gastric H-K-ATPase (Lambrecht et al., 2005). While several KCNE b subunits are found in the central nervous system (CNS), their association with neuronal KV7 channels has not been fully explored and will not be discussed in this review. Neurons are known to express KV7 channels comprised of KV7.2–7.5 a subunits. Some of these gene products may be exclusively neuronal, while others such as KV7.5 can be found in other tissues, such as the skeletal muscle. The first of the central nervous system KV7 genes, KV7.2 and KV7.3 (KCNQ2 and KCNQ3), were discovered using two different approaches. The first approach identified an expressed sequence tag (EST) from public access gene databases that was homologous to the cardiac KVLQT1 (KV7.1; KCNQ1) channel. The EST contained the hallmark pore signature, a GYG amino acid sequence, of Kþ channels and was found only in brain cDNA libraries. Screening of a brain cDNA library with a probe derived from the EST sequence led to the identification of a full-length gene sequence that demonstrated noninactivating Kþ currents upon expression in Xenopus laevis oocytes (Yang et al., 1998). The second approach utilized linkage analysis to identify a chromosomal region in families with inherited epilepsy of newborns and identified the candidate genes by positional cloning. These cloning experiments led to the discovery of two genes; mutations in these genes were responsible for benign familial neonatal convulsions (BFNC). These genes were found to encode KV7 channels and were localized to chromosome 20 for KV7.2 and chromosome 8 for KV7.3 (Biervert et al., 1998; Charlier et al., 1998; Singh et al., 1998). Subsequently, two additional neuronal KV7 family members, KV7.4 (KCNQ4) (Coucke et al., 1999; Kubisch et al., 1999) and KV7.5 (KCNQ5) (Kananura et al., 2000), were identified and cloned. Collectively, these genes were assigned to a subfamily of voltage-gated Kþ channels, the KV7 channel subfamily, by the International Union of Pharmacology (IUPHAR). The original name for these channels was KCNQ, a name assigned by the HUGO Gene Nomenclature Committee (HGNC) (Gutman et al., 2005). In this chapter, as has already been mentioned, we will use the KV7 nomenclature. The chapter will focus only on KV7.2, 7.3, and 7.5 channels. Although KV7.4 is also a neuronal channel, the function and localization of these channels are more restricted. Their most notable known function is their involvement in cochlear hair cell regulation in the inner ear (Coucke et al., 1999; Kubisch et al., 1999; Van et al., 2000; Beisel et al., 2005; Kharkovets et al., 2006). Therefore, this review will be restricted to KV7.2, KV7.3, and,

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to a lesser extent, KV7.5, and they will be referred to collectively as neuronal KV7 channels.

15.2 NEURONAL KV7 CHANNELS: IDENTIFICATION AS M-CHANNELS AND DISTRIBUTION IN THE CENTRAL NERVOUS SYSTEM After the initial cloning, and as studies delineating the localization and expression of the neuronal KV7 channels were published, the potential importance of KV7 channels as regulators of neuronal excitability was immediately apparent, particularly when mutated neuronal KV7 channels, as mentioned above, were shown to cause BFNC (Biervert et al., 1998; Charlier et al., 1998; Singh et al., 1998; Yang et al., 1998; Biervert and Steinlein, 1999). Shortly thereafter, pharmacological and biophysical identity was established between KV7.2/7.3 (and likely KV7.5/7.3) heteromultimers and the elusive “M”-channel (Wang et al., 1998; Brown and Yu, 2000), providing significant new evidence for their importance in neuronal regulation. Native M-channels and the corresponding macroscopic M-current were first characterized in amphibian sympathetic neurons (Brown and Adams, 1980; Adams et al., 1982a). M-channels were notable as they were slowly activating and noninactivating, active at membrane potentials at or near the resting membrane potential of neurons, and muscarinic cholinergic agonists produced a reduction in the Mcurrent, demonstrating a direct and inhibitory link between G-protein-coupled receptors (GPCRs) and a physiological Kþ current. Inhibitory linkages to other receptors were also found, indicating a propensity for channel inhibition by one or more second messenger systems initiated by this class of ligand–receptor interactions (Adams et al., 1982b, 1983 ; Shapiro et al., 1994; Schweitzer, 2000; Wallace et al., 2002; Filippov et al., 2006; Jia et al., 2007). The GPCR-mediated inhibition of neuronal KV7 channels was later found to result from the hydrolysis of membraneassociated phosphatidylinositol 4,5-bisphosphate (PIP2), secondary to the activation of phospholipase C (PLC) (Suh and Hille, 2002; Zhang et al., 2003). This finding has been much extended recently. These data will be discussed briefly here; however, these have been extensively dealt with in Chapter 11 of this book. In the evaluation of emerging drug targets, the putative target’s known contribution to cellular physiology, the distribution of the gene products, and their possible direct linkage with disease via mutation analysis are criteria that add significant weight to the rationale for pursuing a particular small-molecule modulator discovery program. The primary pore-forming components of M-channels, KV7.2, KV7.3, and (probably) KV7.5, are widely distributed in brain and spinal cord. Prior to the discovery of their molecular correlates, M-channel-mediated Kþ currents were found in many neuronal populations by virtue of their characteristic inhibition by activation of certain G-protein-coupled receptors and the relaxation of their current following hyperpolarization. After the cloning of neuronal KV7 channels and their identification as the molecular substrates of M-channels, studies focused on regional gene expression and immunocytochemical analyses of CNS KV7 channel distributions. These studies confirmed the widespread distribution of M-channels, with cortical, thalamic,

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cerebellar, and hippocampal neurons expressing among the highest levels of various combinations of KV7.2, KV7.3, and KV7.5 mRNAs (Lerche et al., 2000; Geiger et al., 2006; Liang et al., 2006; Kanaumi et al., 2007). KV7 mRNA and channel density generally increase in late embryonic and early postnatal development in the rodent and human central nervous system (Smith et al., 2001; Geiger et al., 2006; Kanaumi et al., 2007), after which there may be a regional reduction in channel density. The presence and importance of neuronal KV7/M-channels in regulating neuronal excitability has been well documented in sensory ganglia (Passmore et al., 2003; Rivera-Arconada et al., 2004; Rivera-Arconada and Lopez-Garcia, 2005), an important contributor to their potential utility in pain therapy (see discussion below). In general, the distribution of these channels, both regionally and developmentally, as well as their biophysical characteristics support their role in providing enduring resistance to depolarizing excitatory influences. Mutation analyses demonstrated their involvement in BFNC and suggested their utility as targets for antiepileptic drugs (AEDs) (Rogawski, 2006). Recognition of some forms of chronic pain as a form of neuronal hyperexcitability disorder involving neurons in the sensory/pain pathways (Waxman, 1999), neurons that were known to express KV7 channels as well as other voltage-gated ion channels, suggested that these channels may have value as targets for the discovery of pain therapeutics. Initial studies suggested that the subcellular distribution of KV7 channels was primarily somatodendritic. Recent studies have shown that their subcellular distributions, and therefore the role of the channels, are likely to be more complex. KV7.2/ KV7.3 heteromultimeric M-channels are preferentially localized on the surface of axons and enriched in axon initial segments (Chung et al., 2006; Vervaeke et al., 2006; Yue and Yaari, 2006). Distribution of the channel complex to these loci is differentially regulated. Axon initial segments, an ideal locale for control of spike after hyperpolarization, are mediated by ankyrin-G binding motifs of both KV7.2 and KV7.3 channel subunits, whereas more distal axonal surface expression is regulated by domains within the KV7.2 channel C-terminal region (Chung et al., 2006). Devaux et al. (2004) examined a KV7.2 mutation that produces myokymia and the distribution of KV7.2 and KV7.3 subunits in axons by confocal microscopy. The R207W KV7.2 channel has reduced charge in the S4 voltage sensor (Fig. 15.1a) and shifts the activation voltage curve in the positive (depolarizing) direction (Dedek et al., 2001). Its association with myokymia suggested some form of functional control of motor axon excitability by these channels. Immunocytochemical analyses using polyclonal antibodies to both KV7.2 and KV7.3 channel proteins demonstrated that KV7.2 was found at nodes of Ranvier in these peripheral nerves as well as in the spinal cord and in axon initial segments in much of the CNS (Devaux et al., 2004). KV7.2 and KV7.3, however, were not found to colocalize at these nodes (KV7.3 was primarily found in nodes of smaller diameter fibers), directly demonstrating that while these two channels often form heteromultimeric M-channels, they do not always have overlapping distributions and can have important independent functions. A subsequent study showed that the slow nodal Kþ current IKS, which participates in the pronounced spike-frequency adaptation observed in motor nerve fibers, is mediated exclusively by KV7.2 in large diameter fibers of sensory nerves (Schwarz et al., 2006). Clearly, unlike early simplistic views of

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FIGURE 15.2 (a) Immunocytochemical analysis of KV7.2 protein expression in a dorsal root ganglion. Staining, using an affinity-purified primary antiserum raised against a KV7.2 peptide, reflects the degree of KV7.2 channel expression in neurons of all sizes within the DRG. (b) Staining with a preimmune serum is negative, as is staining in the presence of excess peptide (not shown), demonstrating specificity of the staining shown in panel A.

the function of KV7 channels in neuronal network dynamics, their physiologic roles are complex and only partially understood. For example, it is not known if they have a significant direct role in the regulation of neurotransmitter release at the level of the synaptic bouton, and their role in the regulation of dendritic excitability is also less clear. Our group used in situ hybridization and immunocytochemistry to examine the distribution of KV7.2 channel protein throughout the brain with particular focus on pain pathways. As shown in Fig. 15.2 (Goldstein and Dworetzky, unpublished observations), KV7.2 protein is expressed at highest levels in the large cell and medium cell populations of sensory neurons in the dorsal root ganglion (DRG), with moderate to low levels of expression in the small cell population. Double-label immunofluorescense studies (not shown) demonstrated that some of the large to medium diameter neurons that express the highest levels of KV7.2 protein also express the neuroactive peptide calcitonin gene-related peptide (CGRP). The smaller diameter neurons, which expressed lower levels of KV7.2 protein, all coexpressed substance P. In a study of trigeminal circuitry, RT-PCR techniques applied to dissociated neurons of the trigeminal ganglion gave detectable mRNA signals for KV7.2, KV7.3, and KV7.5 subunits, with the most abundant signal for KV7.2. Further localization studies by in situ hybridization demonstrated the presence of neuronal KV7 channels in the trigeminal nucleus caudalis. Given that the release of CGRP has been closely linked to the onset of migraine pain and both CGRP and substance P have been associated with inflammatory pain (Henry, 1993; Sluka, 1996; Durham, 2006; Doods et al., 2007), modulating KV7 channels in these key regions could significantly impact these pain modalities. These studies are supported by another report examining KV7 members in the nociceptive sensory system (Passmore et al., 2003). Using RT-PCR, they demonstrated the presence of KV7.2, 7.3, and 7.5 mRNA in both small and large DRG neurons. Moreover, confocal microscopy revealed signal in the somata and neuronal

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processes of these neurons (Passmore et al., 2003). An additional localization study has reported the presence of KV7.2, 7.3, and 7.5 in rat visceral sensory neurons isolated from the nodose ganglia (Wladyka and Kunze, 2006). Collectively, these results indicate that neuronal KV7 channels are localized and play an important role in key pain circuitry. It follows from this that openers of these channels could possibly reduce the release of neuroactive peptides involved in pain and/or hyperpolarize these neurons, alleviating either pain initiation and/or transmission of painful stimuli.

15.3 INFLUENCE OF KV7 CHANNELS ON NEURONAL BEHAVIOR Native M-channels and cloned KV7 channels activate at membrane potentials that are close to the resting potential of neurons, unlike most voltage-gated potassium channels. They are relatively slow to activate upon depolarization and slow to deactivate, and, perhaps most important in terms of their functional significance, they are noninactivating. Therefore, in the absence of external modulatory influences, these channels can have enduring effects on membrane excitability within their activation–voltage ranges. Under physiological conditions, as was demonstrated with native M-channels (Brown and Adams, 1980; Constanti and Brown, 1981; Adams et al., 1982a; Selyanko et al., 1992; Stansfeld et al., 1993; Costa and Brown, 1997; Zhu et al., 2000), they can be very effective at regulating the subthreshold excitability of certain neuronal populations, with significant roles in regulating the frequency and ultimately the pattern of action potential discharge in many types of neurons. Their importance in neuronal regulation was punctuated by the discovery that neuronal KV7 mutations lead to BFNC (Biervert et al., 1998; Charlier et al., 1998; Biervert and Steinlein, 1999; Lerche et al., 1999; Hirose et al., 2000; Singh et al., 2003), indicating that reduction or removal of the influence of KV7.2 and KV7.3 channels can dramatically alter neuronal excitability. Most mutations leading to BFNC are autosomal dominant negative mutations, with little or no effect on remaining wild-type KV7 channels. This indicates that the phenotype of BFNC in most cases was caused by a reduction in current, a haploinsufficiency, rather than an alteration in the biophysical characteristics of mutant channels. This was further evidence that neuronal KV7/Mchannels contribute significantly to the maintenance of electrical homeostasis in many neurons, and that modest alteration of the level of expressed current can have profound effects in some systems depending on the stage of development of the organism. As the name implies, BFNC is largely benign and resolves quickly during development. While there remains some increased risk of seizure disorder and even more serious complications in later life, it is clear that the role of KV7 channels is complex and that it changes with development. Members of at least one family with KV7 mutation-related BFNC also have a concomitant form of myokymia (Dedek et al., 2001), an involuntary repetitive series of muscle contractions, indicating additional functions of at least one member of this ion channel subfamily. Knockout animals have not significantly contributed to our understanding of the involvement of these channels in disease states or their potential as targets for the development of treatments due to homozygous lethality or other complicating issues.

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One study disrupted the mouse Kv7.2 gene and heterozygous mutant mice had decreased Kv7.2 expression and demonstrated hypersensitivity to the proconvulsant pentylenetetrazole (Watanabe et al., 2000). Incidentally, there was no mention of a change in nociceptive behavior. Conclusions concerning the physiological roles of neuronal KV7 channels and the importance of development in defining these functions have been greatly strengthened recently by transgenic studies. Peters et al., (2005) used transgenic M-channel-deficient mice conditionally expressing a dominant negative pore mutant construct (hQ2-G279S) in a comprehensive study of the electrophysiological, morphological, developmental, and behavioral consequences of M-channel deficiency. The results were complex; pyramidal cells were lost in mutantmale mice, but therewas muchless cell loss in female mice, demonstrating X-chromosomalinheritance and a different pattern of expression of the mutant channel. These morphological changes in male mice were observed only when the channels were suppressed during early development (approximately from birth to 3 weeks postnatal). While conditional suppression of M-channels following this critical period resulted in altered hippocampal physiology, the morphology was largely unchanged. Under whole-cell voltageclamp, hippocampal pyramidal neurons from male mice expressing the mutant channel (either throughout life or conditionally) had a significant reduction in M-channel current and increased resting input resistance, and responded to depolarizing current pulses with significantly greater numbersofactionpotentials (reduced spike-frequency adaptation), all in keeping with the known functions of these channels. They also had undetectable levels of medium-duration after-hyperpolarizations following action potentials and a reduction in subthreshold electrical resonance behavior in the “theta” frequency range (3–5 Hz). Theta frequency resonance in hippocampal neurons has long been associated with neuronal correlates of learning (Gray and Ball, 1970; Staubli and Xu, 1995; Kinney et al., 1999; McNaughton et al., 2006); and Hu et al. (2002) had recently demonstrated an M-channel contribution to hippocampal theta resonance. This discussion of KV7 channel function is intended to convey some of what may be contributing to the difficulty in readily translating our knowledge of both the physiology and pharmacology of these channels into useful therapeutics. In thinking about the discovery and development of drugs targeting KV7 channels, modulator compounds have usually been thought of in almost a unitary sense, when clearly the channels themselves have multiple and complex roles in many different neurons. In at least two dimensions, they are a multimodal target; they are a subfamily of KV channels, and the same members are likely to have different functions in different neurons and possibly at different loci in the same neurons, and their distributions and functions may change with development. A further layer of complexity possibly affecting the utility and side effect profile of neuronal KV7 modulators is provided by the following discussion of biochemical substrates of KV7 channel regulation.

15.4 REGULATION OF KV7 CHANNELS Transmembrane voltage (Vm) is the primary controller or regulator of neuronal KV7 current. However, well before the molecular entities comprising M-channels had been

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cloned and characterized, this class of Kþ current was known for, and in fact defined by, its modulation by endogenous ligand–receptor interactions (Brown and Adams, 1980; Shapiro et al., 1994; Marrion, 1997). The relatively hyperpolarized voltage– activation ranges of KV7/M-channels means that these channels can contribute significantly to resting levels of neuronal excitability as well as countering some forms of physiologically relevant depolarization. GPCR-mediated or other physiologically relevant inhibition of M-current in neurons produces slow depolarizations, resulting in long-term increases in neuronal excitability, a major contributor to the integration of synaptic influences in many cells. The discovery and the subsequent characterization of the underlying biochemical pathways that connect receptor activation and M-channel inhibition are among the most important and interesting findings associated with these channels. In rat superior cervical ganglion (SCG) neurons, Marrion et al. (1989) demonstrated pharmacologically that muscarinic cholinergic suppression of M-current was probably mediated by receptors of the M1 receptor subtype. Shapiro et al., (2001) later confirmed this, when they found that M1 muscarinic receptor knockout mice lacked muscarinic modulation of M/KV7 channels. The latter group also demonstrated that they could create a functional KV7.2/KV7.3 modulatory system in vitro by coexpressing the channels in tsA-201 cells along with muscarinic M1 receptors, an effect that depended on some unknown intracellular factor but was not a result of intracellular Ca2þ transients (Shapiro et al., 2000, 2001). A diffusible messenger had been implicated in muscarinic suppression of M channels as early as 1992 (Selyanko et al., 1992), and hints of the identity of this factor came from both an increasing awareness of the role of membrane phospholipids in ion channel regulation (Hilgemann et al., 2001), and the earlier work implicating the involvement of phoshotidylinositol 4,5-bisphosphate (PIP2) hydrolysis in bradykinin receptor activation-mediated depolarization of neuroblastoma–glioma hybrid cells, an effect attributable to M-current suppression (Higashida et al., 1986). In 2002, Suh and colleagues used this heterologous coexpression system to examine the recovery of M-current following its suppression by muscarinic cholinergic receptor activation. They found that recovery was dependent on intracellular ATP and was independent of G-protein activity (Suh and Hille, 2002). The need for hydrolysable ATP indicated involvement of one or more kinases in the recovery of native M-current or recombinant KV7-mediated current, so they tested a number of kinase inhibitors in both SCG cells and in the tsA-201 heterologous system. Wortmannin, an inhibitor of Pl 4-kinase (a lipid kinase that, with ATP, replenishes PIP2), as well as other pharmacological inhibitors of PIP2 generation, greatly reduced or eliminated recovery of the current, strongly suggesting PIP2 involvement in M-current recovery (Suh and Hille, 2002). Zhang et al. (2003) directly tested this hypothesis using PIP2 and a number of analogues to determine if PIP2 could both activate M/KV7 channels inhibited by GPCR activation as well as re-activate channels that have run down following insideout patch excision. Rapid channel rundown in excised membrane patches had been a problem since the discovery of M-channels and inhibited single-channel analysis of cloned KV7 channels. Most single-channel work therefore relied on the use of cell-attached recordings, where manipulation of the intracellular and extracellular

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milieu is problematic. Zhang et al. (2003) found that PIP2 and analogues could reestablish current following rundown and the following receptor-mediated inhibition, and that all neuronal KV7 channel subtypes were similarly modulated. These results have been confirmed and extended by several other groups (Post-Munson et al., 2003; Li et al., 2005; Suh et al., 2006; Suh and Hille, 2007). Zhang et al. (2003) also mutated a positively charged amino acid residue in a proximal C-terminal region of KV7.2 that is homologous to PIP2 binding domains in other ion channels; coexpression of these mutated KV7.2 subunits (H328C-KV7.2) with wild-type KV7.3 subunits produced currents of significantly lower amplitude that were much less sensitive to PIP2 and more sensitive to receptor-mediated inhibition. Li et al. (2005) also examined the role of PIP2 in neuronal KV7 channel open probability. While their experiments confirmed the relationship of muscarinic cholinergic suppression of these channels to regulation of PIP2 and the re-activation of rundown channels in excised patches, they also discovered that different subfamily members had different open probabilities when recorded in cell-attached patch configurations (PIP2 system intact). This apparently intrinsic correlation of open probability (at a given voltage) with KV7 subtype was found to correlate with their sensitivity to PIP2, suggesting that open probability is not only a function of the PIP2 available to the channel, but also of the channel’s inherent PIP2 sensitivity. Another study has used palmitoylated peptides (palpeptides) to dynamically probe the intramembrane regulation of neuronal KV7 channels (Robbins et al., 2006). These peptides were designed to interrupt associations such as the coupling of G-proteins to a receptor and receptor activation to PIP2 hydrolysis; their PIP2 palpeptide (palmitoyl-HRQKHFEKRR) was designed based on the assumption that H328 was a C-terminal residue involved in PIP2 binding, as reported previously (Zhang et al., 2003). Robbins and colleagues found that their PIP2 palpeptide rapidly and reversibly reduced current and sensitized currents to muscarinic receptor activation, additional evidence of the importance of PIP2 to neuronal KV7 regulation. It is currently unknown whether PIP2’s regulation of these channels is involved in the activity of small-molecule modulators. We have found that retigabine does not reactivate KV7.2/7.3 heteromultimeric channels after the rundown in inside-out excised membrane patches, but an effect of the compound is usually re-established after PIP2 partially reactivates the channel (Gribkoff, Post-Munson, Boissard, and Dworetzky, in preparation). This suggests that retigabine activation of these channels requires some degree of PIP2 occupancy of its binding site. Intracellular Ca2þ was shown long ago to suppress M-channels (Tokimasa, 1985; Marrion, 1996; Selyanko and Brown, 1996a, 1996b), and Ca2þ/calmodulin (CaM) has been shown to be an important regulator of neuronal KV7 channels (Yus-Najera et al., 2002; Gamper et al., 2005; Shahidullah et al., 2005), with identified binding domains in the C-terminus (Wen and Levitan, 2002; Yus-Najera et al., 2002; Shahidullah et al., 2005). This appears not only to be the case in amphibian neurons and cloned KV7 channels, but also in mammalian central neurons, where it was recently shown that hippocampal neurons expressing a CaM binding motif fusion protein had lower density of M-current; the lower M-current density was associated with a change in neuronal discharge characteristics, including increased action potentials evoked by the depolarizing current and an increase in action potential after depolarizations

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(Shahidullah et al., 2005). These findings are also relevant to receptor-mediated regulation of neuronal KV7 current, since increases in intracellular Ca2þ can be mediated through IP3-coupled receptors, including bradykinin and P2Y receptors, and the effects of activation of these receptors, unlike muscarinic receptors, depend on CaM (Villarroel, 1996; Cruzblanca et al., 1998; Gamper and Shapiro, 2003; Zaika et al., 2007). Gamper et al. (2005) have shown that coexpression of KV7.2, KV7.4, and KV7.5 with CaM produces current suppression, but that coexpression does not suppress KV7.3 (or KV7.1, the cardiac channel). Interestingly, they also found that the KV7 current enhancement produced by the cysteine-modifying reagent N-ethylmaleimide (NEM) was reduced by CaM overexpression, and that the C-terminal domains were involved in the binding/activity of each overlap. Phosphorylation of neuronal KV7 channels via Src tyrosine kinase also produces channel suppression, distinct from GPCR-mediated inhibition (Gamper et al., 2003). Src tyrosine kinase associates with KV7.2–7.5, but appears to actually produce phosphorylation only of KV7.3–7.5; mutation of two tyrosines (tyr-67 and tyr-349) produced additive inhibition (Li et al., 2004). Recently, a physiological role for this additional mechanism of KV7 channel inhibition was demonstrated; activation of epidermal growth factor receptors, a member of the class of receptor tyrosine kinases, was found to produce biphasic inhibition. In a recombinant expression system and in SCG neurons, the faster of the two inhibitory components resulted from PIP2 hydrolysis, while a slower form of inhibition was found to result from tyrosine phosphorylation (Jia et al., 2007). These results serve to indicate the multiple routes of receptor-mediated inhibition of KV7 channel currents, and help to suggest mechanisms of pharmacological modulation that may both explain the actions of known small-molecule modulators as well as suggest additional routes to synthesis of new molecules. Additional phosphorylation sites have recently been identified, one of which is found in the S4–S5 linker region of all neuronal KV7 channels, producing channel inhibition when mutated (Surti et al., 2005). While there are additional mechanisms of neuronal KV7 channel modulation that have recently been elucidated, these examples are sufficient to indicate the importance of understanding the role of biochemical regulation of neuronal KV7 channel function and possibly drug discovery.

15.5 NEURONAL KV7 CHANNEL PHARMACOLOGY 15.5.1

Modulators

Some reference has been made in earlier sections of this chapter to neuronal KV7 channel modulators such as retigabine. At the outset, we think it is important to operationally define some of the terminology used in our description of the pharmacology of these voltage-gated ion channels. Unlike established pharmacological terminology for GPCRs, the mode of action of Kþ channel modulators, in particular, compounds that activate the channel, is still being refined. The application of voltageclamp techniques to the study of ion channel pharmacology enabled detailed biophysical studies of either whole-cell currents or single channels, allowing some

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characterization of the nature of compound–channel interactions, but this has not prevented ongoing confusion around the terminology. The term opener or activator is commonly used throughout the literature, but does not adequately describe the mode of action of all these “positive modulator” compounds. In general, openers or activators are expected to increase the open probability of the channel or increase macroscopic current amplitude, but this nomenclature is really too simplistic. For example, retigabine, the first publicly disclosed KV7 opener, has a complex and interesting profile in that it has inhibitory activity at higher membrane potentials. In our Kþ channel opener programs, we typically found a number of different types of positive modulators in addition to those with the voltage-dependent opener-inhibitor profile of retigabine. As per our experience, neuronal KV7 channel openers may work in concert with the activity of a channel over the “normal” activation–voltage range, enhancing currents without significantly affecting the activation threshold, while others can significantly alter the activation threshold. In addition, some openers appeared to remove the voltage dependence of activation entirely. Whether these effects represent some continuum is currently unclear, since the effects are often concentration dependent. Clearly, the modes of compound interaction that can increase channel current are complex and in most cases not well understood, and the implications of these profiles on neuronal responsiveness and systems physiology remain to be elucidated. Much the same confused terminology concerning compounds that decrease channel current also exists. Pore-blocking peptides, for example, occlude the pore and directly limit ion flux, and are therefore clearly current blockers. However, small molecules that inhibit channel current may not actually block, but rather in many respects act like inverse agonists, reducing current by some means other than directly blocking ion flow through the pore. For the purpose of this study, the term opener is used to describe molecules that increase ion channel current over physiologically relevant levels of Vm, and inhibitor will be used to describe molecules doing the opposite. Almost immediately after the cloning of neuronal Kv7 channels, it became widely accepted that these channels may have significant value as drug targets. The initial publicly disclosed opener of these channels was the antiepileptic drug candidate retigabine (Fig. 15.3) (Main et al., 2000; Wickenden et al., 2000; Schroder et al., 2001; Tatulian et al., 2001), an analogue of the approved pain medication flupirtine (Katadolon) (Mastronardi et al., 1988; Ringe et al., 2003; Boscia et al., 2006). Retigabine is modestly potent and is not highly specific, but it is a very effective opener of KV7.2, KV7.5, and heteromultimeric KV7 channels (Main et al., 2000; Wickenden et al., 2000; Schroder et al., 2001; Tatulian et al., 2001; Tatulian and Brown, 2003; Schenzer et al., 2005). Its effects are characterized by a significant increase in channel current over a narrow voltage range. As mentioned above, at more positive voltages, the opener is less effective, and under some conditions, channel current significantly decreases at more positive voltages relative to control currents (this “crossover” voltage dependence of opener action is a characteristic of many neuronal KV7 channel openers). This effect is also concentration dependent and is more pronounced at higher concentrations. Recent evidence has pinpointed probable sites of interaction of this compound within KV7 channels (Schenzer et al., 2005).

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FIGURE 15.3 Structures of KV7 openers discussed in text, and associated with the experimental or clinical treatment of pain: (a) Marketed compounds, (b) retigabine is in clinical development (the structure of ICA-105665 has not been publicly disclosed), and (c) preclinical discovery compounds used for proof-of-concept studies in animal models.

In 1997, our group initially used the expression of mouse KV7.2 channels, and later human KV7.2/7.3 heteromultimers, to initiate a program targeting these channels as regulators of neuronal excitability. We found that some openers of another important Kþ channel, the large-conductance calcium-activated Kþ (maxi-K, BK, KCa1.1) channel, were also openers of neuronal KV7 channels; this is typified by the fluoroxindole compound BMS-204352, a potent maxi-K channel opener (Gribkoff et al., 2001) whose KV7 channel activity has been cited by others (Schroder et al., 2001, 2003; Dupuis et al., 2002; Jensen, 2002). Over the past 10 years, a number of synthetic programs have been initiated to design neuronal KV7 channel openers (Munro and Dalby-Brown, 2007). It has actually proven quite easy to discover openers of these channels. Openers such as zinc pirithione (Xiong et al., 2007) and the structurally related compounds such as meclofenamic acid and diclofenac have been discovered by academic laboratories (Fig. 15.3) (Peretz et al., 2005). A large number of compounds resulted from our program at Bristol-Myers Squibb (Wu et al., 2003, 2004a, 2004b; Hewawasam et al., 2004; L’Heureux et al., 2005), and additional compounds have been offered or further characterized by teams at Icagen (Wickenden et al., 2008) and NeuroSearch (Schroder et al., 2001, 2003; Dupuis et al., 2002; Jensen, 2002). While other companies have had programs in this area, there has been little public disclosure of specific compounds of interest along with data sufficient to evaluate their potency, efficacy, and specificity. A major struggle with most compounds has been their relative lack of specificity, either between Kþ channel families, as is the case with BMS-204352, or between members of the KV7 subfamily, or both.

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A compound recently reported from the group at Icagen, however, provides some degree of discrimination between neuronal KV7 subfamily members, and reflects their understanding of this issue and their commitment to discovering more subtypespecific modulators. The compound ICA-27243 (N-(6-chloro-pyridin-3-yl)3,4-difluoro-benzamide; Fig. 15.3) (Wickenden et al., 2008) produces a leftward shift in the voltage dependence of KV7.2/7.3 channel activation with an EC50 of 1–2 mM and demonstrates some degree of selectivity, with greater potency and possibly efficacy against KV7.2/7.3 heteromultimeric channels versus either KV7.4 homomultimers or KV7.3/7.5 heteromultimers. In addition, ICA-27243 did not have significant effects on several ion channels and receptors that are the target of current AEDs, such as g-amino butyric acid (GABA) receptors, native high-voltageactivated Ca2þ channels, and a Naþ channel (NaV1.2). As discussed above, in contrast to ICA-27243; retigabine, Bristol-Myers Squibb compounds BMS-204352 and the acrylamide, known publicly as [S]-1 (Fig. 15.3), do not discriminate significantly between subfamily members and appear to require a conserved tryptophan (at position 236 in KV7.2, position 265 in KV7.3, position 242 in KV7.4, and position 235 in KV7.5) in the S5 transmembrane domain to maintain efficacy (Schenzer et al., 2005; Wuttke et al., 2005; Bentzen et al., 2006), as well as possibly other residues in the S6 domain identified as critical for gating (Wuttke et al., 2005). This site of action may not allow big discrimination between neuronal KV7 subfamily members, but it does allow discrimination between neuronal KV7 channels and the cardiac KV7 channel KV7.1, since this site in S5 is not conserved in this subfamily member. Since mutations of this channel (KV7.1) are associated with long QT syndrome (Peroz et al., 2008), its exogenous modulation is to be approached with extreme caution, if at all, and, incidentally, any putative KV7 channel modulator should be assessed for activity at this channel. It is currently unknown whether ICA27243 interacts with the critical tryptophan residue in S5 in neuronal KV7 channels, or whether it has an entirely novel site of interaction that is not conserved between all of the subfamily members. Regardless of the molecular substrate, the compound represents some degree of progress toward subtype specificity and still exhibits activity in models of neuronal hyperexcitability. As mentioned previously, inhibitors of these channels have also been described, most notably linopirdine and XE-991, compounds that originated in a Dupont program for cognition enhancement (Costa and Brown, 1997; Zaczek et al., 1998; Wang et al., 2000). Linopirdine (AVIVA) was the first inhibitor to be tested in clinical trials. The mixed results from these trials and the subsequent removal from further development were attributed to the short and variable half-life in humans and poor brain penetration in animal studies. While additional inhibitors have been discovered in the course of synthetic programs targeting openers, the lack of therapeutic targets for KV7 inhibitors, other than cognition enhancement, has largely relegated these compounds to the status of research tools. This stands in contrast to the situation with openers of neuronal KV7 channels, where localization, functional data, and results of animal studies and, recently, human experimentation have continued to support the idea that the right molecule should have value in several widespread and important neurogenic conditions and diseases, including pain.

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Therapeutic Targets

The spectrum of therapeutic targets for modulators of KV7 channels has not significantly changed since initial descriptions, primarily focusing on chronic pain and epilepsy (Cooper and Jan, 2003; Gribkoff, 2003; Munro and Dalby-Brown, 2007). Some evidence now exists to support anxiety as a target for neuronal KV7 openers (Korsgaard et al., 2005; Hansen et al., 2008), and there is some suggestion that modulators may have utility in the treatment of addiction (Hansen et al., 2007) and dystonia (Richter et al., 2006). In general, these therapeutic targets were determined based on the initial channelopathy (BFNC) findings, the distributions of the channels, the known effects of reference compounds, and the theoretical linkage of disorders other than the epilepsies with selective neuronal “hyperexcitability,” such as migraine pain and other chronic and particularly neuropathic pain (Wall, 1991; Welch et al., 1993; Waxman, 1999). Neuronal KV7 channel inhibitors may have therapeutic value as candidate cognition-enhancing compounds; however, their current value remains as research tools for attenuating or blocking the effects of opener compounds and for pharmacological excision of these currents to study their physiological roles, in particular, neurons and neuronal networks. As mentioned above, the first significant modulators of KV7 channels described publicly were the AED retigabine and the candidate cognition-enhancer compounds linopirdine and XE-991 (Fig 15.3). From the perspective of providing some degree of proof-of-concept in human disease for AEDs (retigabine) and pain (flupirtine, diclofenac), these retrospectively discovered neuronal KV7 modulators have proven useful for increasing confidence in these channels as therapeutic targets.

15.5.3 Neuronal KV7 Modulation and its Effect on Pain Circuitry and Experimental Pain Our direct experience with therapeutic applications for neuronal KV7 modulators has been in the areas of chronic pain and migraine headache. Pain is a broad term used to describe the discomfort, distress, or suffering resulting from the excessive activation of specific subsets of sensory neurons. Acute pain can have significant survival value for an organism by promoting flight or other responses that will minimize further injury. In this vein, acute pain is not usually considered a disorder, although therapeutics directed to acute pain clearly have utility in medical and dental procedures, among others. On the other hand, chronic or semichronic pain syndromes (such as neuropathic pain or cancer pain), or chronic–episodic forms of pain such as migraine can be severe, life altering, and are often of no obvious value to an organism, although in and of themselves they are not really diseases either, but are usually reflections of other conditions. Inflammatory pain occurs after tissue injury and is mediated by the release of multiple proinflammatory factors that result in tissue swelling and the activation of specialized peripheral pain receptors or “nociceptors.” Neuropathic pain often (but not always) results from a lesion or injury to nerves in the peripheral or central nervous system, and is a common and often incapacitating clinical condition largely resistant to treatment with commonly prescribed analgesics. Painful peripheral neuropathies can

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have many etiologies such as trauma and metabolic disorders, but in most cases, there is no known cause. Neuropathic pain is characterized by spontaneous pain, mechanical and thermal hyperalgesia (exaggerated response to a painful stimulus) and allodynia (pain response to an innocuous stimulus). Symptoms are correlated with and may result from the development of “central sensitization” in the spinal cord, which may be initiated by unusually high-frequency and sustained (“ectopic”) spontaneous peripheral nerve activity and a decrease in inhibitory tone; as such it is an example of neuroplasticity run amok (Ossipov et al., 2000; Campbell and Meyer, 2006). In what is no doubt a serious but nevertheless useful oversimplification of pain pathophysiology, central sensitization associated with pain syndromes results from neuronal hyperexcitability and, in turn, produces hyperexcitability in more central components of pain pathways. This phenomenology thus opens the door for treatment of these forms of pain by decreasing this excessive neuronal activity (while hopefully sparing “normal” activity). This logic is identical to that used to ameliorate epileptiform neuronal activity, and suggests that chronic pain can be approached pharmacologically by many of the same drugs such as the AED and neuronal KV7 opener retigabine. There is a growing body of evidence suggesting that neuronal KV7 channels are important regulators of excitability in neurons of pain pathways. For example, the opener retigabine and inhibitors XE-991 and linopirdine were used to investigate the roles of KV7 channels in pain transmission using spinal cord preparations as well as recordings from single spinal neurons. It was shown that motor neurons and neurons in the deep dorsal horn express functional M-currents; application of retigabine produced decreases in the excitability of these neurons, resulting in depression of transmission of nociceptive information, effects that were blocked by XE-991 (Rivera-Arconada et al., 2004; Rivera-Arconada and Lopez-Garcia, 2005, 2006). In another study, both small- and large-diameter-cultured DRG neurons were demonstrated to have M-currents that were enhanced by retigabine and blocked by linopirdine (Passmore et al., 2003). Retigabine was shown to inhibit C and Ad fiber-mediated responses; this effect was blocked by the administration of linopirdine. While the specifics of the neuronal circuitry and neurochemistry underlying pain can be somewhat species specific, acute pain is so important to the survival of organisms that one has to drop to the lowest branches of the phylogenetic tree to convincingly locate animals that do not display behavior reflecting (albeit perhaps somewhat anthropomorphically) acute pain-like responses to potentially harmful stimuli. Chronic pain is somewhat more problematic, but rodents can be shown to respond to some experimental manipulations by the development of exaggerated sensory responses, indicative of the development of chronic neuropathic pain. Some human pain modalities such as migraine, however, remain challenging. Similar to the modeling of psychiatric disorders in animals, some forms of pain like migraine do not have any obvious mechanism for their independent measurement. Nevertheless, in a research setting, both neuropathic pain and inflammatory pain can be modeled in animals for the characterization of pain therapeutics, although there have been some questions concerning the predictive validity of these models (Whiteside et al., 2008). However, with proper care, particularly with respect to the determination of the contribution of side effects (such as lethargy, somnolence, or anesthesia) to apparent

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analgesic effects in particular models with specific compounds, useful and reasonably predictive information can be obtained. Neuropathic pain models range from the use of various types of nerve constrictions or ligations to the use of the antitumor drug streptozotocin to induce diabetes and its attendant neuropathic pain. This latter drug’s toxicity to pancreatic cells is the basis of both its utility in pancreatic cancer and its use in producing experimental diabetes. Changes in pain thresholds due to any of these manipulations are measured in a number of ways. For example, they can be measured mechanically by testing the threshold of tolerated pressure on a paw or the response to thermal pain can be tested by assessing the animal’s tolerance to local changes in skin temperature. Inflammatory pain is induced by injecting (usually) the animal’s paw with formalin, carrageenan, or other chemical stimuli to induce paw flinching, biting, licking, and other responses and the results quantified in terms of an acute response phase, measured within the first 10–15 min, and a persistent response phase, measured between 10–15 min and 90 min after injection. Demonstration of the archetypal broad-spectrum neuronal KV7 channel opener retigabine’s analgesic activity has been reported in several pain models. Retigabine was shown to have analgesic activity in a behavioral model of inflammatory pain produced by intrapaw injection of carrageenan, an effect that was reversed by the application of the inhibitor XE-991 (Passmore et al., 2003). In a spinal nerve ligation model of neuropathic pain in rats, retigabine significantly and dose-dependently increased the pain threshold, an effect that could be diminished with the coadministration of linopirdine (Dost et al., 2004). In a model of neuropathic pain induced by chronic constriction injury as well as in the model of persistent pain induced by formalin, retigabine demonstrated efficacy under some test conditions (BlackburnMunro and Jensen, 2003). With respect to modeling different types of pain and understanding the broad utility of neuronal KV7 modulation, visceral sensory neurons of the nodose ganglion have been shown to express KV7.2/7.3/7.5 proteins, and M-currents could be recorded from these cells. Flupirtine, an analogue of retigabine approved in Europe for the treatment of pain, increased M-current and hyperpolarized neuronal resting membrane potentials in these cells in culture (Wladyka and Kunze, 2006). Consistent with this, in a capsaicin-induced visceral pain model, retigabine dose-dependently suppressed visceral pain behavior (Hirano et al., 2007). In a study of pain of musculoskeletal origin, retigabine produced antinociceptive effects in a model of acid-induced muscle allodynia (Nielsen et al., 2004). Taken together, these data demonstrate a high degree of proof-of-concept for the role of neuronal KV7 channel modulation in the treatment of various neuropathic and inflammatory pain conditions in animals. At the Bristol-Myers Squibb Company, we had an extensive program targeting pain by modulating neuronal KV7 channels. Many of these compounds proved to be analgesic at doses that produced no significant effects on independent measures of general motor disruption and somnolence. One such compound, the cinamamide (S)-3(2-fluoro- phenyl)-N-(1-[3-(pyridine-3-yloxy)-phenyl]-ethyl)-acrylamide (Fig. 15.3), was tested in several pain models using i.v. administration. In the formalin test of inflammatory pain, the compound produced a significant reduction in both the early and late phases of the response profile using flinching as the measure of pain response

NEURONAL KV7 POTASSIUM CHANNELS AS EMERGING TARGETS

Threshold (g)

410

**

**

15

** ** **

10

Gabapentin (100 mg/kg)

* BMS-Cinnamamide ex. (10 mg/kg)

5 PEG400 (0.5 mL/kg) i.v.

0 –15 0

30

60

90

Time (min)

FIGURE 15.4 Effects of a neuronal KV7 opener in a rat model of neuropathic pain. The BMScinnamamide induced a significant attenuation of neuropathic pain behavior at 10 mg/kg i.v., peaking 15 min after injection. A lesser but significant effect was also observed at 3 mg/kg i.v. (not shown). This particular cinnamamide has a modest duration of action due to either rapid metabolism or high clearance. Gabapentin (100 mg/kg i.v.) has a slower onset of action but longer duration of effect. (n ¼ 8–10 rats per group. *P < 0.05, **P < 0.01 versus PEG-400, Dunnett’s test).

(not shown). In a spinal nerve ligation model, the compound significantly reduced tactile allodynia, although its duration of action was limited (Fig. 15.4). The compound was also active in the streptozotocin model of diabetic neuropathy (not shown). Retigabine, like the Bristol-Myers Squibb compounds discussed above (and below for migraine), is a broad-spectrum opener of neuronal KV7 channels. Retigabine also interacts with other potential pharmacological targets. Collectively, these compounds leave room for improvement in understanding the potential contribution of neuronal KV7 channels in general and specific subfamily members in particular to analgesia related to their activation. The Icagen compound introduced previously (ICA-27243; Fig. 15.3) is a much more discriminating opener of KV7.2/7.3 channels relative to its effects on other subfamily members, and was reported to show efficacy in a wide range of anticonvulsant models (Wickenden et al., 2008). ICA-27243 also demonstrated analgesic efficacy in the rat carrageenan model of inflammatory pain, the rat formalin model of chronic pain, and also antihyperalgesia and antiallodynic efficacy in the rat chronic constrictive injury model of neuropathic pain. Icagen has recently advanced a compound, ICA-105665 (the structure has not been disclosed), into phase 1 clinical trials to assess its safety and pharmacokinetic profiles, with a stated goal of advancing the compound into clinical trials as an AED and as a pain therapeutic (www.Icagen.com). As already mentioned, the use of KV7 openers for the treatment of migraine pain is another interesting therapeutic indication for these compounds. In part, this was based on the localization of these channels within neuronal populations associated with migraine headache (such as the trigeminal nuclei) and on the concept that this condition falls within the broad spectrum of “hyperexcitability disorders.” Migraine is characterized by unilateral throbbing head pain that is often associated with visual disturbances and is considered to be a chronic or chronic–episodic disorder. It has been suggested that cortical spreading depression may underlie various prodromes

SUMMARY AND DISCUSSION

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that precede the onset of migraine headache, particularly visual aura (Lauritzen, 1994). Cortical spreading depression is defined as a wave of neuronal excitation followed by long-lasting inhibition that spreads from a focal point. Clinical neurological migraine prodromes proceed in a temporal fashion that is correlated with the expected rate of spreading depression. These neurological symptoms, with associated changes in blood flow, correlate well with spreading depression phenomena (Lauritzen, 1994). More recently, spreading depression was visualized during triggered migraine attacks in humans using functional magnetic resonance imaging (Cao et al., 1999). This evidence suggests that spreading depression may underlie the visual aura and possibly other prodromes that precede migraine and cause the ensuing migraine attack and accompanying pain (Hardebo, 1992). Similar to the modeling of neuropathic and inflammatory pain for the testing of preclinical drug candidates, it is possible to produce cortical spreading depression in laboratory animals. In view of these findings, attenuation of cortical spreading depression induced by placing crystals of KCl onto the cortex has become a currently accepted practice to evaluate and compare novel compounds for their potential in the prophylactic treatment of migraine headache. We standardized this model using valproic acid, a drug sometimes used for the prophylactic treatment of migraine headache. Using the BMS-acrylamide (S)-1 KV7 opener (Fig. 15.3), we found that the compound produced a reduction in the total number of cortical spreading depressions induced by the KCl. The potency compared to valproic acid was greater, while efficacy was reduced relative to this reference compound, thus providing additional support for the broad utility of this target in the treatment of pain (Wu et al., 2003).

15.6 SUMMARY AND DISCUSSION We have presented a review of previous work, including our own contributions, in which we characterize this new subfamily of important voltage-dependent Kþ channels and introduce their pharmacology. We believe that these channels represent interesting targets for the development of important new therapeutics in a number of areas, and we have most closely focused on their potential utility in the treatment of serious and chronic pain syndromes. There remains a large unmet medical need for the treatment of pain, and developing compounds against new targets is a good strategy for attempting to circumvent the shortcomings of existing medications. Many lines of evidence support the hypothesis that neuronal KV7 channel openers would be useful for the treatment of different types of pain, particularly chronic neuropathic pain and migraine. The neuronal KV7 channels are localized in key neuronal circuitry involved in pain pathways; the key role the M-current plays in controlling cell firing is well documented and preclinical proof-of-concept studies demonstrate their efficacy in a wide array of pain models with at least three different neuronal KV7 channel openers. This is an area that is currently seeing significant activity, including clinical trials of at least one opener of these channels. Valeant pharmaceuticals is currently testing retigabine in phase 3 trials for the adjunctive treatment for partial-onset seizures after it

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was demonstrated to be efficacious in reducing monthly seizure rates in a phase 2 trial (Porter et al., 2007). In addition, Valeant has filed an IND application for retigabine (as an immediate release formulation) in the treatment of pain associated with postherpetic neuralgia (www.Valeant.com). In addition, both flupirtine and diclofenac are effective and approved for use in human pain (Mastronardi et al., 1988; Ringe et al., 2003; Moore, 2007). None of these compounds are specific, however, they can have a number of side effects, depending on the dose. Most important, their other pharmacology may very well contribute as much or more to their efficacy in pain as their ability to open KV7 channels (among other things diclofenac is a COX inhibitor and flupirtine is an NMDA receptor antagonist). It is unclear whether new molecules such as ICA-27243 represent a significant advance in terms of therapeutic potential, as opposed to the demonstrated progress in molecular pharmacology. Determining this will require comprehensive analyses of new compounds’ effects in animal models of disease, compared to activity profiles in models predictive of dose-limiting side effects. Finally, we will have to compare these profiles with those obtained under similar conditions with other molecules not displaying this degree of subfamily and molecular target specificity. To approach pain as it has been approached with sodium and calcium channels, additional key data will be required. For example, we know that alternative splice variants exist for neuronal KV7 channel members (Smith et al., 2001), however, there has been little or no work to determine if there is any functional (modal) significance to their expression. Specifically, are there defined sensory neuronal populations expressing specific KV7.2–7.5 variants, which could be specifically targeted for pain, as may be the case for N-type (CaV2.2) Ca2þ channels (see Chapter 8) (Bell et al., 2004; Altier et al., 2007)? Also, are channel splice variants sufficiently distinct to provide a substrate for differential pharmacology, something that has not yet been convincingly or usefully shown for other voltage-gated ion channels? Are there functional neuronal KCNE beta subunits, and are they involved in changing the biophysical and pharmacological characteristics of neuronal KV7 channels as they are with peripheral KV7.1 channels? Most important, is there something about the voltage dependence of KV7 modulators, a feature that is readily apparent in the actions of many openers, which can be exploited to provide state dependency of drug action in a mode favoring therapeutic utility? Drug discovery efforts directed toward detection and exploitation of these more subtle aspects of drug action have greater promise to produce effective pain medications with minimal side effects.

REFERENCES Adams PR, Brown DA, Constanti A, 1982a. M-currents and other potassium currents in bullfrog sympathetic neurones. J Physiol 330: 537–572. Adams PR, Brown DA, Constanti A, 1982b. Pharmacological inhibition of the M-current. J Physiol 332: 223–262. Adams PR, Brown DA, Jones SW, 1983. Substance P inhibits the M-current in bullfrog sympathetic neurones. Br J Pharmacol 79: 330–333.

REFERENCES

413

Altier C, Dale CS, Kisilevsky AE, Chapman K, Castiglioni AJ, Matthews EA, Evans RM, Dickenson AH, Lipscombe D, Vergnolle N, Zamponi GW, 2007. Differential role of N-type calcium channel splice isoforms in pain. J Neurosci 27: 6363–6373. Barhanin J, Lesage F, Guillemare E, Fink M, Lazdunski M, Romey G, 1996. KvLQT1 and lsK (minK) proteins associate to form the Iks cardiac potassium current. Nature 384: 78–80. Beisel KW, Rocha-Sanchez SM, Morris KA, Nie L, Feng F, Kachar B, Yamoah EN, Fritzsch B, 2005. Differential expression of KCNQ4 in inner hair cells and sensory neurons is the basis of progressive high-frequency hearing loss. J Neurosci 25: 9285–9293. Bell TJ, Thaler C, Castiglioni AJ, Helton TD, Lipscombe D, 2004. Cell-specific alternative splicing increases calcium channel current density in the pain pathway. Neuron 41: 127– 138. Bentzen BH, Schmitt N, Calloe K, Dalby BW, Grunnet M, Olesen SP, 2006. The acrylamide (S)-1 differentially affects Kv7 (KCNQ) potassium channels. Neuropharmacology 51: 1068–1077. Biervert C, Steinlein OK, 1999. Structural and mutational analysis of KCNQ2, the major gene locus for benign familial neonatal convulsions. Hum Genet 104: 234–240. Biervert C, Schroeder BC, Kubisch C, Berkovic SF, Propping P, Jentsch TJ, Steinlein OK, 1998. A potassium channel mutation in neonatal human epilepsy. Science 279: 403–406. Blackburn-Munro G, Jensen BS, 2003. The anticonvulsant retigabine attenuates nociceptive behaviours in rat models of persistent and neuropathic pain. Eur J Pharmacol 460: 109–116. Boscia F, Annunziato L, Taglialatela M, 2006. Retigabine and flupirtine exert neuroprotective actions in organotypic hippocampal cultures. Neuropharmacology 51: 283–294. Brown DA, Adams PR, 1980. Muscarinic suppression of a novel voltage-sensitive Kþ current in a vertebrate neurone. Nature 283: 673–676. Brown BS, Yu SP, 2000. Modulation and genetic identification of the M channel. Prog Biophys Mol Biol 73: 135–166. Campbell JN, Meyer RA, 2006. Mechanisms of neuropathic pain. Neuron 52: 77–92. Cao Y, Welch KM, Aurora S, Vikingstad EM, 1999. Functional MRI-BOLD of visually triggered headache in patients with migraine. Arch Neurol 56: 548–554. Charlier C, Singh NA, Ryan SG, Lewis TB, Reus BE, Leach RJ, Leppert M, 1998. A pore mutation in a novel KQT-like potassium channel gene in an idiopathic epilepsy family. Nat Genet 18: 53–55. Chung HJ, Jan YN, Jan LY, 2006. Polarized axonal surface expression of neuronal KCNQ channels is mediated by multiple signals in the KCNQ2 and KCNQ3 C-terminal domains. Proc Natl Acad Sci USA 103: 8870–8875. Constanti A, Brown DA, 1981. M-currents in voltage-clamped mammalian sympathetic neurones. Neurosci Lett 24: 289–294. Cooper EC, Jan LY, 2003. M-channels: neurological diseases, neuromodulation, and drug development. Arch Neurol 60: 496–500. Costa AM, Brown BS, 1997. Inhibition of M-current in cultured rat superior cervical ganglia by linopirdine: mechanism of action studies. Neuropharmacology 36: 1747–1753. Coucke PJ, Van HP, Kelley PM, Kunst H, Schatteman I, Van VD, Meyers J, Ensink RJ, Verstreken M, Declau F, Marres H, Kastury K, Bhasin S, McGuirt WT, Smith RJ, Cremers CW, Van de HP, Willems PJ, Smith SD, Van CG, 1999. Mutations in the KCNQ4 gene are responsible for autosomal dominant deafness in four DFNA2 families. Hum Mol Genet 8: 1321–1328.

414

NEURONAL KV7 POTASSIUM CHANNELS AS EMERGING TARGETS

Cruzblanca H, Koh DS, Hille B, 1998. Bradykinin inhibits M current via phospholipase C and Ca2þ release from IP3-sensitive Ca2þ stores in rat sympathetic neurons. Proc Natl Acad Sci USA 95: 7151–7156. Dedek K, Kunath B, Kananura C, Reuner U, Jentsch TJ, Steinlein OK, 2001. Myokymia and neonatal epilepsy caused by a mutation in the voltage sensor of the KCNQ2 Kþ channel. Proc Natl Acad Sci USA 98: 12272–12277. Devaux JJ, Kleopa KA, Cooper EC, Scherer SS, 2004. KCNQ2 is a nodal Kþ channel. J Neurosci 24: 1236–1244. Doods H, Arndt K, Rudolf K, Just S, 2007. CGRP antagonists: unravelling the role of CGRP in migraine. Trends Pharmacol Sci 28: 580–587. Dost R, Rostock A, Rundfeldt C, 2004. The anti-hyperalgesic activity of retigabine is mediated by KCNQ potassium channel activation. Naunyn Schmiedebergs Arch Pharmacol 369: 382–390. Dupuis DS, Schroder RL, Jespersen T, Christensen JK, Christophersen P, Jensen BS, Olesen SP, 2002. Activation of KCNQ5 channels stably expressed in HEK293 cells by BMS-204352. Eur J Pharmacol 437: 129–137. Durham PL, 2006. Calcitonin gene-related peptide (CGRP) and migraine. Headache 46 (Suppl 1): S3–S8. Filippov AK, Choi RC, Simon J, Barnard EA, Brown DA, 2006. Activation of P2Y1 nucleotide receptors induces inhibition of the M-type Kþ current in rat hippocampal pyramidal neurons. J Neurosci 26: 9340–9348. Gamper N, Shapiro MS, 2003. Calmodulin mediates Ca2þ-dependent modulation of M-type Kþ channels. J Gen Physiol 122: 17–31. Gamper N, Stockand JD, Shapiro MS, 2003. Subunit-specific modulation of KCNQ potassium channels by Src tyrosine kinase. J Neurosci 23: 84–95. Gamper N, Li Y, Shapiro MS, 2005. Structural requirements for differential sensitivity of KCNQ Kþ channels to modulation by Ca2þ/calmodulin. Mol Biol Cell 16: 3538–3551. Geiger J, Weber YG, Landwehrmeyer B, Sommer C, Lerche H, 2006. Immunohistochemical analysis of KCNQ3 potassium channels in mouse brain. Neurosci Lett 400: 101–104. Gray JA, Ball GG, 1970. Frequency-specific relation between hippocampal theta rhythm, behavior, and amobarbital action. Science 168: 1246–1248. Gribkoff VK, 2003. The therapeutic potential of neuronal KCNQ channel modulators. Expert Opin Ther Targets 7: 737–748. Gribkoff VK, Starrett JE Jr, Dworetzky SI, Hewawasam P, Boissard CG, Cook DA, Frantz SW, Heman K, Hibbard JR, Huston K, Johnson G, Krishnan BS, Kinney GG, Lombardo LA, Meanwell NA, Molinoff PB, Myers RA, Moon SL, Ortiz A, Pajor L, Pieschl RL, PostMunson DJ, Signor LJ, Srinivas N, Taber MT, Thalody G, Trojnacki JT, Wiener H, Yeleswaram K, Yeola SW, 2001. Targeting acute ischemic stroke with a calcium-sensitive opener of maxi-K potassium channels. Nat Med 7: 471–477. Gutman GA, Chandy KG, Grissmer S, Lazdunski M, Mckinnon D, Pardo LA, Robertson GA, Rudy B, Sanguinetti MC, Stuhmer W, Wang X, 2005. International Union of Pharmacology LIII. nomenclature and molecular relationships of voltage-gated potassium channels. Pharmacol Rev 57: 473–508. Hansen HH, Andreasen JT, Weikop P, Mirza N, Scheel-Kruger J, Mikkelsen JD, 2007. The neuronal KCNQ channel opener retigabine inhibits locomotor activity and reduces forebrain excitatory responses to the psychostimulants cocaine, methylphenidate and phencyclidine. Eur J Pharmacol 570: 77–88.

REFERENCES

415

Hansen HH, Waroux O, Seutin V, Jentsch TJ, Aznar S, Mikkelsen JD, 2008. Kv7 channels: interaction with dopaminergic and serotonergic neurotransmission in the CNS. J Physiol. Hardebo JE, 1992. A cortical excitatory wave may cause both the aura and the headache of migraine. Cephalalgia 12: 75–80. Henry JL, 1993. Substance P and inflammatory pain: potential of substance P antagonists as analgesics. Agents Actions Suppl 41: 75–87. Hewawasam P, Chen N, Ding M, Natale JT, Boissard CG, Yeola S, Gribkoff VK, Starrett J, Dworetzky SI, 2004. The synthesis and structure–activity relationships of 3-amino-4benzylquinolin-2-ones: discovery of novel KCNQ2 channel openers. Bioorg Med Chem Lett 14: 1615–1618. Higashida H, Streaty RA, Klee W, Nirenberg M, 1986. Bradykinin-activated transmembrane signals are coupled via No or Ni to production of inositol 1,4,5-trisphosphate, a second messenger in NG108-15 neuroblastoma-glioma hybrid cells. Proc Natl Acad Sci USA 83: 942–946. Hilgemann DW, Feng S, Nasuhoglu C, 2001. The complex and intriguing lives of PIP2 with ion channels and transporters. Sci STKE 2001: RE19. Hirano K, Kuratani K, Fujiyoshi M, Tashiro N, Hayashi E, Kinoshita M, 2007. Kv7.2–7.5 voltage-gated potassium channel (KCNQ2-5) opener, retigabine reduces capsaicin-induced visceral pain in mice. Neurosci Lett 413: 159–162. Hirose S, Zenri F, Akiyoshi H, Fukuma G, Iwata H, Inoue T, Yonetani M, Tsutsumi M, Muranaka H, Kurokawa T, Hanai T, Wada K, Kaneko S, Mitsudome A, 2000. A novel mutation of KCNQ3 (c 925T ! C) in a Japanese family with benign familial neonatal convulsions. Ann Neurol 47: 822–826. Howard RJ, Clark KA, Holton JM, Minor DL Jr, 2007. Structural insight into KCNQ Kv7 channel assembly and channelopathy. Neuron 53: 663–675. Hu H, Vervaeke K, Storm JF, 2002. Two forms of electrical resonance at theta frequencies, generated by M-current, h-current and persistent Naþ current in rat hippocampal pyramidal cells. J Physiol 545: 783–805. Jensen BS, 2002. BMS-204352: a potassium channel opener developed for the treatment of stroke. CNS Drug Rev 8: 353–360. Jia Q, Jia Z, Zhao Z, Liu B, Liang H, Zhang H, 2007. Activation of epidermal growth factor receptor inhibits KCNQ2/3 current through two distinct pathways: membrane PtdIns(4,5)P2 hydrolysis and channel phosphorylation. J Neurosci 27: 2503–2512. Kananura C, Biervert C, Hechenberger M, Engels H, Steinlein OK, 2000. The new voltagegated potassium channel KCNQ5 and neonatal convulsions. Neuroreport 11: 2063– 2067. Kanaumi T, Takashima S, Iwasaki H, Itoh M, Mitsudome A, Hirose S, 2007. Developmental changes in KCNQ2 and KCNQ3 expression in human brain: possible contribution to the age-dependent etiology of benign familial neonatal convulsions. Brain Dev . Kharkovets T, Dedek K, Maier H, Schweizer M, Khimich D, Nouvian R, Vardanyan V, Leuwer R, Moser T, Jentsch TJ, 2006. Mice with altered KCNQ4 Kþ channels implicate sensory outer hair cells in human progressive deafness. EMBO J 25: 642–652. Kinney GG, Patino P, Mermet-Bouvier Y, Starrett JE Jr, Gribkoff VK, 1999. Cognitionenhancing drugs increase stimulated hippocampal theta rhythm amplitude in the urethaneanesthetized rat. J Pharmacol Exp Ther 291: 99–106.

416

NEURONAL KV7 POTASSIUM CHANNELS AS EMERGING TARGETS

Korsgaard MP, Hartz BP, Brown WD, Ahring PK, Strobaek D, Mirza NR, 2005. Anxiolytic effects of Maxipost (BMS-204352) and retigabine via activation of neuronal Kv7 channels. J Pharmacol Exp Ther 314: 282–292. Kubisch C, Schroeder BC, Friedrich T, Lutjohann B, El-Amraoui A, Marlin S, Petit C, Jentsch TJ, 1999. KCNQ4, a novel potassium channel expressed in sensory outer hair cells, is mutated in dominant deafness. Cell 96: 437–446. L’Heureux A, Martel A, He H, Chen J, Sun LQ, Starrett JE, Natale J, Dworetzky SI, Knox RJ, Harden DG, Weaver D, Thompson MW, Wu YJ, 2005. (SE)-N-[1-(3-heteroarylphenyl) ethyl]-3-(2-fluorophenyl)acrylamides: synthesis and KCNQ2 potassium channel opener activity. Bioorg Med Chem Lett 15: 363–366. Lambrecht NW, Yakubov I, Scott D, Sachs G, 2005. Identification of the K efflux channel coupled to the gastric H-K-ATPase during acid secretion. Physiol Genomics 21: 81–91. Lauritzen M, 1994. Pathophysiology of the migraine aura. The spreading depression theory. Brain 117 (Part 1): 199–210. Lerche H, Biervert C, Alekov AK, Schleithoff L, Lindner M, Klinger W, Bretschneider F, Mitrovic N, Jurkat-Rott K, Bode H, Lehmann-Horn F, Steinlein OK, 1999. A reduced Kþ current due to a novel mutation in KCNQ2 causes neonatal convulsions. Ann Neurol 46: 305–312. Lerche C, Scherer CR, Seebohm G, Derst C, Wei AD, Busch AE, Steinmeyer K, 2000. Molecular cloning and functional expression of KCNQ5, a potassium channel subunit that may contribute to neuronal M-current diversity. J Biol Chem 275: 22395–22400. Li Y, Langlais P, Gamper N, Liu F, Shapiro MS, 2004. Dual phosphorylations underlie modulation of unitary KCNQ Kþ channels by Src tyrosine kinase. J Biol Chem 279: 45399–45407. Li Y, Gamper N, Hilgemann DW, Shapiro MS, 2005. Regulation of Kv7 (KCNQ) Kþ channel open probability by phosphatidylinositol 4,5-bisphosphate. J Neurosci 25: 9825–9835. Liang GH, Jin Z, Ulfendahl M, Jarlebark L, 2006. Molecular analyses of KCNQ1-5 potassium channel mRNAs in rat and guinea pig inner ears: expression, cloning, and alternative splicing. Acta Otolaryngol 126: 346–352. Main MJ, Cryan JE, Dupere JR, Cox B, Clare JJ, Burbidge SA, 2000. Modulation of KCNQ2/ 3 potassium channels by the novel anticonvulsant retigabine. Mol Pharmacol 58: 253–262. Marrion NV, 1996. Calcineurin regulates M channel modal gating in sympathetic neurons. Neuron 16: 163–173. Marrion NV, 1997. Control of M-current. Annu Rev Physiol 59: 483–504. Marrion NV, Smart TG, Marsh SJ, Brown DA, 1989. Muscarinic suppression of the M-current in the rat sympathetic ganglion is mediated by receptors of the M1-subtype. Br J Pharmacol 98: 557–573. Mastronardi P, D’Onofrio M, Scanni E, Pinto M, Frontespezi S, Ceccarelli MG, Bianchi F, Mazzarella B, 1988. Analgesic activity of flupirtine maleate: a controlled double-blind study with diclofenac sodium in orthopaedics. J Int Med Res 16: 338–348. McNaughton N, Ruan M, Woodnorth MA, 2006. Restoring theta-like rhythmicity in rats restores initial learning in the Morris water maze. Hippocampus 16: 1102–1110. Moore N, 2007. Diclofenac potassium 12.5 mg tablets for mild to moderate pain and fever: a review of its pharmacology, clinical efficacy and safety. Clin Drug Investig 27: 163–195.

REFERENCES

417

Munro G, Dalby-Brown W, 2007. Kv7 (KCNQ) channel modulators and neuropathic pain. J Med Chem 50: 2576–2582. Nielsen AN, Mathiesen C, Blackburn-Munro G, 2004. Pharmacological characterisation of acid-induced muscle allodynia in rats. Eur J Pharmacol 487: 93–103. Ossipov MH, Lai J, Malan TP Jr, Porreca F, 2000. Spinal and supraspinal mechanisms of neuropathic pain. Ann N Y Acad Sci 909: 12–24. Passmore GM, Selyanko AA, Mistry M, Al-Qatari M, Marsh SJ, Matthews EA, Dickenson AH, Brown TA, Burbidge SA, Main M, Brown DA, 2003. KCNQ/M currents in sensory neurons significance for pain therapy. J Neurosci 23: 7227–7236. Peretz A, Degani N, Nachman R, Uziyel Y, Gibor G, Shabat D, Attali B, 2005. Meclofenamic acid and diclofenac, novel templates of KCNQ2/Q3 potassium channel openers, depress cortical neuron activity and exhibit anticonvulsant properties. Mol Pharmacol 67: 1053–1066. Peroz D, Rodriguez N, Choveau F, Baro I, Merot J, Loussouarn G, 2008. Kv7.1 (KCNQ1) properties and channelopathies. J Physiol. Peters HC, Hu H, Pongs O, Storm JF, Isbrandt D, 2005. Conditional transgenic suppression of M channels in mouse brain reveals functions in neuronal excitability, resonance and behavior. Nat Neurosci 8: 51–60. Porter RJ, Partiot A, Sachdeo R, Nohria V, Alves WM, 2007. Randomized, multicenter, doseranging trial of retigabine for partial-onset seizures. Neurology 68: 1197–1204. Post-Munson DJ, Dworetzky SI, Thompson MW, Gribkoff VK, 2003. A synthetic PIP2 short chain analog modulates cloned hKCNQ2/3 channels. Society for Neuroscience Abstracts on-line (www.sfn.org). Richter A, SanderSE,Rundfeldt C, 2006. Antidystonic effects ofKv7 (KCNQ) channel openers in the dt sz mutant, an animal model of primary paroxysmal dystonia. Br J Pharmacol 149: 747–753. Ringe JD, Miethe D, Pittrow D, Wegscheider K, 2003. Analgesic efficacy of flupirtine in primary care of patients with osteoporosis-related pain. A multivariate analysis Arzneimittelforschung 53: 496–502. Rivera-Arconada I, Lopez-Garcia JA, 2005. Effects of M-current modulators on the excitability of immature rat spinal sensory and motor neurones. Eur J Neurosci 22: 3091–3098. Rivera-Arconada I, Lopez-Garcia JA, 2006. Retigabine-induced population primary afferent hyperpolarisation in vitro. Neuropharmacology 51: 756–763. Rivera-Arconada I, Martinez-Gomez J, Lopez-Garcia JA, 2004. M-current modulators alter rat spinal nociceptive transmission: an electrophysiological study in vitro. Neuropharmacology 46: 598–606. Robbins J, Marsh SJ, Brown DA, 2006. Probing the regulation of M(Kv7) potassium channels in intact neurons with membrane-targeted peptides. J Neurosci 26: 7950–7961. Rogawski MA, 2006. Diverse mechanisms of antiepileptic drugs in the development pipeline. Epilepsy Res 69: 273–294. Sanguinetti MC, Curran ME, Zou A, Shen J, Spector PS, Atkinson DL, Keating MT, 1996. Coassembly of KvLQT1 and minK ( IsK) proteins to form cardiac IKs potassium channel. Nature 384: 80–83. Schenzer A, Friedrich T, Pusch M, Saftig P, Jentsch TJ, Grotzinger J, Schwake M, 2005. Molecular determinants of KCNQ Kv7 Kþ channel sensitivity to the anticonvulsant retigabine. J Neurosci 25: 5051–5060. Schroeder BC, Waldegger S, Fehr S, Bleich M, Warth R, Greger R, Jentsch TJ, 2000. A constitutively open potassium channel formed by KCNQ1 and KCNE3. Nature 403: 196–199.

418

NEURONAL KV7 POTASSIUM CHANNELS AS EMERGING TARGETS

Schroder RL, Jespersen T, Christophersen P, Strobaek D, Jensen BS, Olesen SP, 2001. KCNQ4 channel activation by BMS-204352 and retigabine. Neuropharmacology 40: 888–898. Schroder RL, Strobaek D, Olesen SP, Christophersen P, 2003. Voltage-independent KCNQ4 currents induced by (þ/
)BMS-204352. Pflugers Arch 446: 607–616. Schwarz JR, Glassmeier G, Cooper EC, Kao TC, Nodera H, Tabuena D, Kaji R, Bostock H, 2006. KCNQ channels mediate Iks, a slow Kþ current regulating excitability in the rat node of Ranvier. J Physiol 573: 17–34. Schweitzer P, 2000. Cannabinoids decrease the Kþ M-current in hippocampal CA1 neurons. J Neurosci 20: 51–58. Selyanko AA, Brown DA, 1996a. Intracellular calcium directly inhibits potassium M channels in excised membrane patches from rat sympathetic neurons. Neuron 16: 151–162. Selyanko AA, Brown DA, 1996b. Regulation of M-type potassium channels in mammalian sympathetic neurons: action of intracellular calcium on single channel currents. Neuropharmacology 35: 933–947. Selyanko AA, Stansfeld CE, Brown DA, 1992. Closure of potassium M-channels by muscarinic acetylcholine-receptor stimulants requires a diffusible messenger. Proc Biol Sci 250: 119–125. Shahidullah M, Santarelli LC, Wen H, Levitan IB, 2005. Expression of a calmodulin-binding KCNQ2 potassium channel fragment modulates neuronal M-current and membrane excitability. Proc Natl Acad Sci USA 102: 16454–16459. Shapiro MS, Wollmuth LP, Hille B, 1994. Angiotensin II inhibits calcium and M current channels in rat sympathetic neurons via G proteins. Neuron 12: 1319–1329. Shapiro MS, Roche JP, Kaftan EJ, Cruzblanca H, Mackie K, Hille B, 2000. Reconstitution of muscarinic modulation of the KCNQ2/KCNQ3 Kþ channels that underlie the neuronal M current. J Neurosci 20: 1710–1721. Shapiro MS, Gomeza J, Hamilton SE, Hille B, Loose MD, Nathanson NM, Roche JP, Wess J, 2001. Identification of subtypes of muscarinic receptors that regulate Ca2þ and Kþ channel activity in sympathetic neurons. Life Sci 68: 2481–2487. Singh NA, Charlier C, Stauffer D, DuPont BR, Leach RJ, Melis R, Ronen GM, Bjerre I, Quattlebaum T, Murphy JV, McHarg ML, Gagnon D, Rosales TO, Peiffer A, Anderson VE, Leppert M, 1998. A novel potassium channel gene, KCNQ2, is mutated in an inherited epilepsy of newborns. Nat Genet 18: 25–29. Singh NA, Westenskow P, Charlier C, Pappas C, Leslie J, Dillon J, Anderson VE, Sanguinetti MC, Leppert MF, 2003. KCNQ2 and KCNQ3 potassium channel genes in benign familial neonatal convulsions: expansion of the functional and mutation spectrum. Brain 126: 2726– 2737. Sluka KA, 1996. Pain mechanisms involved in musculoskeletal disorders. J Orthop Sports Phys Ther 24: 240–254. Smith JS, Iannotti CA, Dargis P, Christian EP, Aiyar J, 2001. Differential expression of kcnq2 splice variants: implications to m current function during neuronal development. J Neurosci 21: 1096–1103. Stansfeld CE, Marsh SJ, Gibb AJ, Brown DA, 1993. Identification of M-channels in outside-out patches excised from sympathetic ganglion cells. Neuron 10: 639–654. Staubli U, Xu FB, 1995. Effects of 5-HT3 receptor antagonism on hippocampal theta rhythm, memory, and LTP induction in the freely moving rat. J Neurosci 15: 2445–2452.

REFERENCES

419

Suh BC, Hille B, 2002. Recovery from muscarinic modulation of M current channels requires phosphatidylinositol 4,5-bisphosphate synthesis. Neuron 35: 507–520. Suh BC, Hille B, 2007. Regulation of KCNQ channels by manipulation of phosphoinositides. J Physiol 582: 911–916. Suh BC, Inoue T, Meyer T, Hille B, 2006. Rapid chemically induced changes of PtdIns(4,5)P2 gate KCNQ ion channels. Science 314: 1454–1457. Surti TS, Huang L, Jan YN, Jan LY, Cooper EC, 2005. Identification by mass spectrometry and functional characterization of two phosphorylation sites of KCNQ2/KCNQ3 channels. Proc Natl Acad Sci USA 102: 17828–17833. Tatulian L, Brown DA, 2003. Effect of the KCNQ potassium channel opener retigabine on single KCNQ2/3 channels expressed in CHO cells. J Physiol 549: 57–63. Tatulian L, Delmas P, Abogadie FC, Brown DA, 2001. Activation of expressed KCNQ potassium currents and native neuronal M-type potassium currents by the anti-convulsant drug retigabine. J Neurosci 21: 5535–5545. Tokimasa T, 1985. Intracellular Ca2þ-ions inactivate Kþ-current in bullfrog sympathetic neurons. Brain Res 337: 386–391. Van HP, Coucke PJ, Ensink RJ, Huygen P, Cremers CW, Van CG, 2000. Mutations in the KCNQ4 Kþ channel gene, responsible for autosomal dominant hearing loss, cluster in the channel pore region. Am J Med Genet 93: 184–187. Vervaeke K, Gu N, Agdestein C, Hu H, Storm JF, 2006. Kv7/KCNQ/M-channels in rat glutamatergic hippocampal axons and their role in regulation of excitability and transmitter release. J Physiol 576: 235–256. Villarroel A, 1996. M-current suppression in PC12 cells by bradykinin is mediated by a pertussis toxin-insensitive G-protein and modulated by intracellular calcium. Brain Res 740: 227–233. Wall PD, 1991. Neuropathic pain and injured nerve: central mechanisms. Br Med Bull 47: 631– 643. Wallace DJ, Chen C, Marley PD, 2002. Histamine promotes excitability in bovine adrenal chromaffin cells by inhibiting an M-current. J Physiol 540: 921–939. Wang HS, Pan Z, Shi W, Brown BS, Wymore RS, Cohen IS, Dixon JE, McKinnon D, 1998. KCNQ2 and KCNQ3 potassium channel subunits: molecular correlates of the M-channel. Science 282: 1890–1893. Wang HS, Brown BS, McKinnon D, Cohen IS, 2000. Molecular basis for differential sensitivity of KCNQ and Iks channels to the cognitive enhancer XE991. Mol Pharmacol 57: 1218– 1223. Watanabe H, Nagata E, Kosakai A, Nakamura M, Yokoyama M, Tanaka K, Sasai H, 2000. Disruption of the epilepsy KCNQ2 gene results in neural hyperexcitability. J Neurochem 75: 28–33. Waxman SG, 1999. The molecular pathophysiology of pain: abnormal expression of sodium channel genes and its contributions to hyperexcitability of primary sensory neurons. Pain Suppl 6: S133–S140. Wehling C, Beimgraben C, Gelhaus C, Friedrich T, Saftig P, Grotzinger J, Schwake M, 2007. Self-assembly of the isolated KCNQ2 subunit interaction domain. FEBS Lett 581: 1594– 1598. Welch KM, Barkley GL, Tepley N, Ramadan NM, 1993. Central neurogenic mechanisms of migraine. Neurology 43: S21–S25.

420

NEURONAL KV7 POTASSIUM CHANNELS AS EMERGING TARGETS

Wen H, Levitan IB, 2002. Calmodulin is an auxiliary subunit of KCNQ2/3 potassium channels. J Neurosci 22: 7991–8001. Whiteside GT, Adedoyin A, Leventhal L, 2008. Predictive validity of animal pain models? A comparison of the pharmacokinetic–pharmacodynamic relationship for pain drugs in rats and humans. Neuropharmacology. 54: 767–775. Wickenden AD, Yu W, Zou A, Jegla T, Wagoner PK, 2000. Retigabine, a novel anti-convulsant, enhances activation of KCNQ2/Q3 potassium channels. Mol Pharmacol 58: 591–600. Wickenden AD, Krajewski JL, London B, Wagoner PK, Wilson WA, Clark S, Roeloffs R, NaughtonSmith G, Rigdon GC, 2008. N-(6-chloro-pyridin-3-yl)-3,4 difluoro-benzamide: ICA-27243: a novel, selective KCNQ2/Q3 potassium channel activator. Mol Pharmacol. 73: 977–986. Wladyka CL, Kunze DL, 2006. KCNQ/M-currents contribute to the resting membrane potential in rat visceral sensory neurons. J Physiol 575: 175–189. Wu YJ, Boissard CG, Greco C, Gribkoff VK, Harden DG, He H, L’Heureux A, Kang SH, Kinney GG, Knox RJ, Natale J, Newton AE, Lehtinen-Oboma S, Sinz MW, Sivarao DV, Starrett JE Jr, Sun LQ, Tertyshnikova S, Thompson MW, Weaver D, Wong HS, Zhang L, Dworetzky SI, 2003. (S)-N-[1-(3-morpholin-4-ylphenyl)ethyl]- 3-phenylacrylamide an orally bioavailable KCNQ2 opener with significant activity in a cortical spreading depression model of migraine. J Med Chem 46: 3197–3200. Wu YJ, Boissard CG, Chen J, Fitzpatrick W, Gao Q, Gribkoff VK, Harden DG, He H, Knox RJ, Natale J, Pieschl RL, Starrett JE Jr, Sun LQ, Thompson M, Weaver D, Wu D, Dworetzky SI, 2004a. (S)-N-[1-(4-cyclopropylmethyl-3,4-dihydro-2H-benzo[1,4]oxazin-6-yl)-ethyl]-3-(2fluoro-phenyl)-acrylamide is a potent and efficacious KCNQ2 opener which inhibits induced hyperexcitability of rat hippocampal neurons. Bioorg Med Chem Lett 14: 1991–1995. Wu YJ, Sun LQ, He H, Chen J, Starrett JE Jr, Dextraze P, Daris JP, Boissard CG, Pieschl RL, Gribkoff VK, Natale J, Knox RJ, Harden DG, Thompson MW, Fitzpatrick W, Weaver D, Wu D, Gao Q, Dworetzky SI, 2004b. Synthesis and KCNQ2 opener activity of N-(1benzo[1,3]dioxol-5-yl-ethyl, N-[1-(2,3-dihydro-benzofuran-5-yl)-ethyl, and N-[1-(2,3dihydro-1H-indol-5-yl)-ethyl acrylamides. Bioorg Med Chem Lett 14: 4533–4537. Wuttke TV, Seebohm G, Bail S, Maljevic S, Lerche H, 2005. The new anticonvulsant retigabine favors voltage-dependent opening of the Kv7.2 (KCNQ2) channel by binding to its activation gate. Mol Pharmacol 67: 1009–1017. Xiong Q, Sun H, Li M, 2007. Zinc pyrithione-mediated activation of voltage-gated KCNQ potassium channels rescues epileptogenic mutants. Nat Chem Biol 3: 287–296. Yang WP, Levesque PC, Little WA, Conder ML, Ramakrishnan P, Neubauer MG, Blanar MA, 1998. Functional expression of two KvLQT1-related potassium channels responsible for an inherited idiopathic epilepsy. J Biol Chem 273: 19419–19423. Yue C, Yaari Y, 2006. Axo-somatic and apical dendritic Kv7/M channels differentially regulate the intrinsic excitability of adult rat CA1 pyramidal cells. J Neurophysiol 95: 3480–3495. Yus-Najera E, Santana-Castro I, Villarroel A, 2002. The identification and characterization of a noncontinuous calmodulin-binding site in noninactivating voltage-dependent KCNQ potassium channels. J Biol Chem 277: 28545–28553. Zaczek R, Chorvat RJ, Saye JA, Pierdomenico ME, Maciag CM, Logue AR, Fisher BN, Rominger DH, Earl RA, 1998. Two new potent neurotransmitter release enhancers, 10,10-bis(4-pyridinylmethyl)-9(10H)-anthracenone and 10,10-bis(2-fluoro-4-pyridinylmethyl)-9(10H)-anthracenone: comparison to linopirdine. J Pharmacol Exp Ther 285: 724–730.

REFERENCES

421

Zaika O, Tolstykh GP, Jaffe DB, Shapiro MS, 2007. Inositol triphosphate-mediated Ca2þ signals direct purinergic P2Y receptor regulation of neuronal ion channels. J Neurosci 27: 8914–8926. Zhang H, Craciun LC, Mirshahi T, Rohacs T, Lopes CM, Jin T, Logothetis DE, 2003. PIP(2) activates KCNQ channels, and its hydrolysis underlies receptor-mediated inhibition of M currents. Neuron 37: 963–975. Zhu G, Okada M, Murakami T, Kamata A, Kawata Y, Wada K, Kaneko S, 2000. Dysfunction of M-channel enhances propagation of neuronal excitability in rat hippocampus monitored by multielectrode dish and microdialysis systems. Neurosci Lett 294: 53–57.

16 SMALL-MOLECULE MODULATORS OF LARGE-CONDUCTANCE, CALCIUM-ACTIVATED (BK) CHANNELS John E. Starrett Jr. Discovery Chemistry, Bristol-Myers Squibb Co., 5 Research Parkway, Wallingford, CT 06492, USA

16.1 INTRODUCTION Large-conductance, calcium-activated potassium (maxi-K, BKCa, big K, or BK) channels are present in a wide variety of excitable and nonexcitable cells, including neurons (Meech, 1978), pancreas (Findlay et al., 1985), skeletal muscles (Lerche et al., 1995), and mammalian smooth muscles such as trachea (McCann and Welsh, 1986), colon (Carl et al., 1992), and bladder (Suarez-Kurtz et al., 1991; Zografos et al., 1992). They are also found within the vasculature, including coronary (Sargent et al., 1993) and cerebral vessels (Holland et al., 1996). BK channels have single-channel conductance values in excess of 100 pS, yet maintain a high degree of selectivity for potassium (Latorre et al., 1989). Because of their widespread tissue distribution and dependence on voltage and calcium for activation, BK channel regulation represents an attractive approach to treat a number of diseases associated with abnormalities of smooth muscle and neurons. Due to their large conductance and gating by the critical intracellular messenger calcium, modulation of relatively few BK channels can have profound effects on cell and tissue functions. As knowledge of

Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

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BK channel regulation by small molecules is growing, so is the number of reports of the utility of those small molecules. In model systems, openers have demonstrated activity for the treatment of smooth muscle disorders associated with erectile dysfunction, urinary incontinence, irritable bowel syndrome, and hypertension. BK blockers may be of utility for the treatment of glaucoma and malignant glioma. This review will focus on small-molecule regulators of BK channels that are reported to directly modulate the BK channel. There are also reports of molecules that regulate BK channels via second messenger cascades, such as those induced by stimulation of canabinoid, protinoid, and kinase receptors, which may impact diseases such as neurogenic inflammation of the airway, hypertension, stroke, and glaucoma; however, these approaches are outside the scope of the current study. There have been several reviews focusing on small-molecule regulators of BK channels (Starrett et al., 1996; Gribkoff et al., 1997; Calderone, 2002; Nardi et al., 2003); the current study will concentrate on small-molecule BK channel modulators that have been reported since 2002.

16.2 BK CHANNEL OPENERS Ohwada et al. (2003) have studied the effects of analogues of dehydroabietic acid (Fig. 16.1a) for their ability to open cloned maxi-K channels in human embryonic kidney (HEK) 293 cells. Introducing chlorines at the 12 and 14 positions provided the analogue depicted in Fig. 16.1b, which significantly increased channel-opening activity. The carboxylic acid appears to play an important role for activity in this class of molecules; reduction of the acid to afford the alcohol, depicted in Fig. 16.1c, resulted in complete loss of activity. A more detailed examination of the properties of 12,14-dichloro-dehydroabietic acid (Fig. 16.1b) revealed that it is a potent opener of BK channel a subunits expressed in HEK cells, as measured in inside-out patches via changing the voltage and calcium sensitivity of the channel (Sakamoto et al., 2006). It has an inverse voltage dependence for BK channel activation, but may not have a significant interaction with the BKb1 subunit. The potentiation of BK channels was significantly larger at negative potentials as well as at lower calcium concentrations.

FIGURE 16.1

The dehydroabietic acids and alcohol openers of BK channels.

BK CHANNEL OPENERS

FIGURE 16.2

425

Benzimidazolone and diphenyl hetereocyclic BK openers.

A series of diphenyl-substituted heterocycles were synthesized and evaluated by electrophysiological techniques as openers of cloned BK channels expressed in Xenopus oocytes (Romine et al., 2002). Deannulation of the prototypical benzimidazolones NS-004 and NS-1619, shown in Fig. 16.2a and b, respectively, afforded derivatives such as the trifluoromethylphenyl oxadiazole, shown in Fig. 16.2c, which opened BK channels to 138% of control when evaluated at 20 mM. For comparison, under these conditions, NS-004 and NS-1619 opened BK channels to 132% and 116% of control, respectively. Decreasing the electron density of the phenyl ring by the addition of another trifluoromethyl group afforded the structure depicted in Fig. 16.2d, which opened BK channels to 159% of control. The structure–activity relationship that emerged from this study indicated a lipophilic pocket exists adjacent to the binding region of the channel. The presence of a hydrogen bond donor on the heterocyclic ring was found to be of paramount importance for BK channel activity in these series of compounds. For example, the triazole (Fig. 16.2e) was devoid of activity. The presence of an alcohol capable of hydrogen bonding to a carbonyl is a recurring theme in a variety of BK openers (Starrett et al., 1996). Calderone and coworkers have published a series of papers examining the vasodilatory activity of a series of bis-aryl heterocycles and bis-aryl amides. In these papers, compounds were evaluated for their ability to relax thoracic aortic rings of male normotensive rats precontracted with potassium chloride. Electrophysiologic data were not presented, but the authors speculated that the relaxant properties of these compounds are mediated through BK channels by structural homology to known BK openers such as NS-1619 and the triazolone shown in Fig. 16.2d. In a series of

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FIGURE 16.3 Bis-phenyl amide and triazole BK openers.

bis-phenyl amides, the phenol shown in Fig. 16.3a demonstrated a pIC50 ¼ 7.56 and an efficacy (Emax %) ¼ 86 (Biagi et al., 2004b). Examination of a series of 1,2,3 triazoles revealed the ester illustrated in Fig. 16.3b as the most potent analogue, with a pIC50 ¼ 6.55 and an efficacy (Emax %) ¼ 56 (Biagi et al., 2004a). By comparison, NS-1619 had a pIC50 ¼ 5.18 and an efficacy (Emax %) ¼ 100. Extension of the 1,2,3 triazoles to bis-benzylated versions (Fig. 16.3c) resulted in a phenol with a pIC50 ¼ 7.56 and an efficacy (Emax %) ¼ 95 (Calderone et al., 2005). Replacement of a phenyl group in the bis-phenyl amide series uncovered a pyridine and thiophene, as shown in Fig. 16.3d and e, respectively, as the most active analogues, with full efficacy (100%) and pIC50 ¼ 4.93 for both compounds (Calderone et al., 2006b). Similar to previous reports, replacement of the phenol with a methoxy group resulted in a marked decrease in efficacy and potency. Reversing the position of the amide functionality of bis-phenyl amides (Fig. 16.3a) provided the chlorophenol, shown in Fig. 16.3f, with pIC50 ¼ 6.16 and full efficacy (Calderone et al., 2006a). The plant-derived sapogenin, diosgenin, depicted in Fig. 16.4, increased the activity of BK channels in human cortical neurons (HCN-1A cells) when applied to the external surface of the cells, with an EC50 ¼ 25 mM (Wang et al., 2006). The increased current amplitude occurred without altering the single-channel conductance. Application of diosgenin to the intracellular surface had no effect on BK channels.

16.3 CENTRAL NERVOUS SYSTEM (CNS) THERAPEUTICS 16.3.1

Stroke/Traumatic Brain Injury

Following a stroke or traumatic brain injury, neurons may undergo membrane depolarization, leading to increased intracellular calcium levels and neurotoxicity.

CENTRAL NERVOUS SYSTEM (CNS) THERAPEUTICS

FIGURE 16.4

427

The BK opener diosgenin.

Opening BK channels will lead to membrane hyperpolarization, blocking the further opening of voltage-activated calcium channels. Employing pharmacologic activators of BK channels may activate them at lower levels of intracellular calcium and membrane depolarization, thereby providing a neuroprotective effect. A series of 4-phenyl-3-aminoquinolin-2ones derivatives were synthesized and evaluated as activators of cloned BK channels express in Xenopus oocytes (Hewawasam et al., 2002a). The most active compound to emerge from this study was the trifluoromethyl sulfonamide, shown in Fig. 16.5a, which opened channels to 343% of control at the test concentration of 20 mM. The acidity of the proton plays an important role in the activity of the molecule, as indicated by the weaker increases in current exhibited by the methylsulfonamide, shown in Fig. 16.5b, and the acetamide, shown in Fig. 16.5c (164% and 120% of control, respectively). For

FIGURE 16.5 4-Phenyl-3-amino quinolinone BK openers.

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SMALL-MOLECULE MODULATORS

the purpose of treating stroke, the sulfonamide depicted in Fig. 16.5a could not be advanced further due to undetectable brain levels of the compound following i.v. administration of 5 mg/kg to rats. The parent 3-aminoquinolinone (Fig. 16.5d), which opened BK channels to 186% of control, was determined to achieve brain levels of 1.4 mM and a brain/plasma ratio of 5.3 at 2 h following i.v. administration. To determine the ability of the parent 3-aminoquinolinone to reduce cell loss resulting from neuronal ischemia, a standard rodent model of focal ischemia was employed, which involved permanent occlusion of middle cerebral artery in spontaneously hypertensive rats (MCAO model). When administered i.v. 30 min postMCAO at a dose of 1 mg/kg, it induced a significant reduction (14%, p < 0.05) in neocortical infarct volume. A series of 1,3,4 oxadiazoles were studied in Xenopus oocytes as BK openers (Romine et al., 2007). Similar to the bis-aryl heterocycles discussed above, deannulation of NS-004 and optimization led to BMS-191011, as shown in Fig. 16.6a. When evaluated at 1 mM, BMS-191011 opened cloned BK channels to 126% of control. In rats, BMS-191011 demonstrated a brain/plasma ratio of 21 at 15 min following i.v. administration. In a permanent occlusion stroke model, BMS-191011 produced a significant reduction in infarct volume when administered 2 h following occlusion of the middle cerebral artery in rats. One shortcoming of BMS-191011 was the low aqueous solubility (25 mg/mL. Incubation in rat and human plasma indicated cleavage of the prodrug to afford BMS-191011 and the prodrug was also cleaved in rats following i.v. administration to provide BMS191011 in blood and brain. After stirring for 24 h in PEG400/water, >97% of the prodrug remained intact. When evaluated in a permanent occlusion stroke model, the prodrug significantly reduced cortical infarct volume following i.v. administration to rats.

FIGURE 16.6 BMS-191011.

The BK channel openers BMS-191011 and BMS-204352, and a prodrug of

CENTRAL NERVOUS SYSTEM (CNS) THERAPEUTICS

429

The fluorooxindole BMS-204352 (Maxipost, Fig 16.6c) opens cloned BK channels to 247% of control at 10 mM in HEK 293 cells (Hewawasam et al., 2002b). In a rat permanent occlusion model of stroke, BMS-204352 significantly reduced infarct volume when administered 2 h following the occlusion (Gribkoff et al., 2001). In a rat model of traumatic brain injury, administration of 0.1 mg/kg BMS204352 improved neurological motor function 1 and 2 weeks postinjury and reduced the extent of cerebral edema in the ipsilateral hippocampus, thalamus, and adjacent cortex (Cheney et al., 2001). Administration of 0.03 mg/kg of BMS-204352 reduced cerebral edema in the ipsilateral thalamus. BMS-204352 underwent a series of clinical trials (the potassium-channel opener stroke trials; POST), including two phase III efficacy trials, which were multicentered, worldwide clinical trials, enrolling 1500 patients (Starrett et al., 2004). The drug was administered at doses of 1 mg/day or 0.1 mg/day for 4 days following a stroke. The trials were methodologically sound and showed a positive effect for TPA; however, BMS-204352 was not statistically different from placebo. Due to the low aqueous solubility of BMS-204352, a Tween-80 formulation was employed to administer the drug intravenously. Dose escalation in POST was limiting because the formulation became hypotensive. It was concluded that alternative formulations or a prodrug would be needed to safely administer higher doses to adequately test the hypothesis of utilizing a BK opener as a neuroprotectant following a stroke. Cilostazol (Fig. 16.7a) has been approved in the United States as a cyclic nucleotide phosphodiesterase 3 inhibitor for reduction in intermittent claudation. By elevating cyclic AMP, it acts as a vasodilator and antiplatelet aggregator (Hong et al., 2006). In patients with recurrent cerebral infarction, cilostazol demonstrated a reduced risk of stroke. Subsequent to the clinical findings, cilostazol was found to have BK channel opening properties (Hong et al., 2003). In SK–N–SH cells, cilostazol opened BK channels in a concentration-dependent manner as determined in whole-cell patches. The opening could be blocked by the BK channel blocker IbTx, but not by the KATP blocker glibenclamide. In a rat model of ischemia, cilostazol significantly reduced infarct volume when administered 1 h after the completion of a 2 h occlusion of the middle cerebral artery. In in vitro studies, cilostazol, as well as BMS-204352,

FIGURE 16.7

The BK channel openers cilostazol, KR 31378, and zonisamide.

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SMALL-MOLECULE MODULATORS

prevented a TNF-a-induced decrease in viability with suppressed DNA fragmentation, along with an increase in CK2 phosphorylation and a decrease in PTEN phosphorylation. A downstream effect of this BK-mediated second messenger system was an increase in Akt/CREB phosphorylation, increase in Bcl-2 protein, and decrease in Bax protein, leading to prevention of cell death. The benzopyran KR 31378, illustrated in Fig. 16.7b, has also demonstrated BKopening and anti-apoptotic properties via a similar pathway as cilostazol (Kim et al., 2004). When administered to rats i.p. at 30 or 50 mg/kg at 5 min, 4 h, and 8 h after the completion of 2 h of ischemia of the middle cerebral artery, KR 31378 demonstrated a significant reduction in cerebral infarct as well as a reduction in TUNEL-positive cells, a decrease in DNA fragmentation, an increase in Bcl-2, and decreases in Bax and cytochrome C expression (Hong et al., 2002). 16.3.2

Epilepsy/Bipolar Disorder

The benzisoxazole sulfonamide zonisamide, illustrated in Fig. 16.7c, is approved in the United States for the treatment of epilepsy. The mechanism of action of zonisamide is not well understood, but there are reports that it blocks voltage-dependent sodium channels and T-type calcium channels. A recent report showed that clinically relevant brain concentrations (30 mM) of zonisamide can activate BK channels (Huang et al., 2007). In neuron-derived H19-7 cells, zonisamide increased the activity of BK channels in a concentration-dependent manner by significantly increasing the channel open probability. The authors speculate that it is reasonable to assume that BK channels present in neurons are relevant targets of the action of zonisamide. In addition to its action as an anticonvulsant, zonisamide has also demonstrated clinical efficacy in bipolar disorder (Ghaemi et al., 2006). In an 8-week study conducted in bipolar outpatients, a significant improvement in the depressive symptoms was observed.

16.4 SMOOTH MUSCLE THERAPEUTICS 16.4.1

Erectile Dysfunction (ED)

Among the various types of potassium channels present in corporal smooth muscle, BK channels are the most prominent subtype. Opening of BK channels in corporal smooth muscle may lead to membrane hyperpolarization, resulting in smooth muscle relaxation, an increase in corporal blood flow, and an improvement in erection function. Several different groups have studied the effects of BK channels and their role in ED. Hewawasam et al. (2003b) have described several series of 4-aryl quinolinones demonstrating in vivo activity in a rat model of erectile dysfunction. The compounds were initially characterized for their ability to increase potassium-mediated wholecell outward currents using two-electrode voltage-clamp recording from Xenopus oocytes expressing cloned hSlo BK channels. Structure–activity relationship studies were conducted in a series of 3-aryl-6-trifluoromethyl quinolinones, depicted in

SMOOTH MUSCLE THERAPEUTICS

FIGURE 16.8

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Quinolinone BK channel openers.

Fig. 16.8a, in which the C-3 alkoxy substituent (R2) was optimized. In all examples, the phenol (R1 ¼ H) was significantly more active than the corresponding methyl ethers (R1 ¼ CH3). The most effective opener identified in this series was ethyl alcohol BMS223131, shown in Fig. 16.8b. At the test concentration of 20 mM, it opened BK channels to 378% of control. In phenylepherine-contracted rabbit cavernosum strips, BMS-223131 produced an 83% inhibition of isomeric force. The relaxation of rabbit cavernosum strips induced by BMS-223131 could be reversed by iberiotoxin (IbTX), a selective BK channel blocker, indicating that the tissue relaxation induced by BMS223131 was mediated through the opening of BK channels. In an in vivo rat model of erectile function, BMS-223131 was administered i.v. at a dose of 1 mg/kg. It had no effect on basal intracavernous pressure, but produced a significant potentiation of electrically evoked increases in intracavernous pressure 50 min postdose, indicating that BK openers such as BMS-223131 may be of value in treating humans with erectile dysfunction. Further evaluation of BMS-223131 revealed that it may exist as two stable atropisomers (Vrudhula et al., 2007). Separation of BMS-223131 by chiral highperformance liquid chromatography (HPLC) resulted in isolation of (þ)-BMS223131 and (
)-BMS-223131, whose absolute stereochemistry was determined by single X-ray crystallography. Electrophysiological evaluation of the atropisomers determined the (
) and (þ) isomers that opened cloned BK channels to 292 and 214% of control, respectively, at the test concentration of 10 mM. At this concentration, racemic BMS-223131 opened BK channels to 259% of control. In an effort to extend the activity observed in alcohol BMS-223131, a series of 3-quinolinyl mercaptans were studied (Hewawasam et al., 2004). The amino

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SMALL-MOLECULE MODULATORS

alcohol, shown in Fig. 16.8c, opened BK channels to 192% of control in Xenopus oocytes expressing hSlo. It also demonstrated a considerable improvement in aqueous solubility compared to related analogues, with water solubility of >1.9 mg/mL. At the test concentration of 10 mM, it inhibited 55% of isometric force in isolated rabbit corpus cavernosum precontracted with phenylepherine; at a concentration of 1 mg/kg i.v., it produced a significant increase in electrically stimulated intracavernous pressure in a rat model of erectile function. Further work in the 3-quinolinyl mercaptans by Boy et al. (2004) identified compounds extremely efficacious at opening cloned BK channels expressed in oocytes. The N-methyl piperazine, depicted in Fig. 16.8d, opened BK channels to 474% of control at the test concentration of 20 mM and the N,N-diethyl amine, shown in Fig. 16.8e, opened channels to 192% of control. Although numerous efficacious openers were identified in this study, in many cases the opening did not translate into relaxation of phenylepherine-contracted rabbit corpus cavernosum strips. The authors speculate that this disconnect in activity may be due to differences in tissue penetration issues, or may be due to differences in the channels expressed in oocytes versus those expressed in cavernosum strips. The most efficacious compound in the rabbit strips in this series was the amine, shown in Fig. 16.8e, which produced a 57% relaxation of rabbit cavernosum strips at the test concentration of 10 mM. Studies conducted by Nelson and coworkers have examined the effects of BK channels on erectile function in BK channel knockout mice (Slo
/
) (Werner et al., 2005). In cavernosum tissue from (Slo
/
) mice, precontracted with phenylepherine, a fourfold increase in phasic contractions was observed. In electrically stimulated cavernosum tissue from (Slo
/
) mice, relaxation was reduced to 50%, similar to the reduction observed in wild-type cavernosum tissue treated with the BK blocker iberiotoxin. Intracavernous pressure in vivo exhibited significant oscillations in (Slo
/
) mice, but not wild-type mice. Electrically stimulated increases in intracavernous pressure were reduced by 22% in (Slo
/
) mice. The authors strongly support the idea that the BK channel could be an important target in treating patients with erectile dysfunction. Melman et al. (2006) have taken the concept of treating erectile dysfunction by opening BK channels into the clinic by treating patients with injections of hMaxiK, a naked DNA plasmid carrying the human cDNA encoding hSlo. A gene transfer approach that overexpresses BK channels in the corpus cavernosum may overcome malfunctioning or underexpressed endogenous BK channels in men with ED. In a phase I study, 11 patients were given a single-dose caverosum injection of hMaxi-K. The injection was well tolerated with no adverse events or clinical laboratory abnormalities. This trial was not designed to study efficacy; however, one patient in each of the high-dose groups reported clinically significant and sustained improvement in erectile function. The authors suggest that further clinical trials should be conducted with larger patient groups to confirm the safety and efficacy of this approach. The gene transfer therapy approach may also be applicable to other smooth muscle disorders such as urinary incontinence, irritable bowel syndrome, and asthma.

SMOOTH MUSCLE THERAPEUTICS

16.4.2

433

Irritable Bowel Syndrome

BK channel openers may also be of utility for the treatment of irritable bowel syndrome. BK channels are expressed in the gastrointestinal track where they influence the action potentials of the vagal motor neurons (Pedarzani et al., 2000) as well as primary afferent neurons responsible for normalizing stretch-induced excitation (Kunze et al., 2000) and secretomotor reflexes (Clerc and Furness, 2002). Opening of BK channels may therefore result in an increase in transit time in the gut. In addition to demonstrating activity in a rat model of erectile dysfunction (discussed above), BMS-223131 (Fig. 16.8b) has also shown activity in a rat model of irritable bowel syndrome (Sivarao et al., 2005). Following i.p. administration of 3, 6, and 20 mg/kg, BMS-223131 markedly inhibited fecal output in response to stress as well as demonstrated a significant reduction in moisture content. A reduction in stressinduced colonic motility as well as in visceral nociception was also observed. 16.4.3

Urinary Incontinence

Indoles, as shown in Fig. 16.9 are claimed to open BK channels in freshly isolated bladder smooth muscle cells as well as relax isolated rabbit urinary bladder strips precontracted with potassium chloride (Lee et al., 2005). Specific data were not provided, but the inventors claim that the compounds in the patent application are useful for the treatment of urinary incontinence or overactive bladder. 16.4.4

General Smooth Muscle Relaxants

Examination of a series of derivatives of ketoconazole, as shown in Fig. 16.10a, for their ability to open BK channels expressed in bovine smooth muscle cells has identified single variations on the scaffold which changed the characteristics from a BK channel blocker to an opener (Power et al., 2006). The bromophenyl analogue, depicted in Fig. 16.10b, produced a 95% blockade of BK currents at 30 mM, whereas the aminophenyl analogue, shown in Fig. 16.10c, produced a 566% opening at the same concentration. The imidazole is not required for opening activity as demonstrated by the 235% opening of the racemic mixture of 1:1 cis:trans isomers of the methyl analogue, illustrated in Fig. 16.10d. Mallotoxin (Fig. 16.11a; rottlerin; MTX) is a natural product capable of opening BK channels, with the unusual property of opening BK channels in a voltagedependent and calcium-independent manner (Zakharov et al., 2005). Mallotoxin has been reported to inhibit protein kinase C d with an IC50 ¼ 3
6 mM; however,

FIGURE 16.9

An indole BK channel opener.

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SMALL-MOLECULE MODULATORS

FIGURE 16.10 The BK channel modulators ketoconazole and derivatives of ketoconazole.

activation of BK channels by mallotoxin was observed at 100 nM. As a potential smooth muscle relaxant, this study relates the folk remedy of kamala for the treatment of tapeworm. Mallotoxin is purified from kamala, suggesting that the antihelmenthic effects may be a result of BK channel activation by mallotoxin, relaxing smooth muscle in the tapeworm, resulting in a decrease in egg-laying behavior. The authors suggest that the Mallotoxin scaffold may be utilized to enhance channel activity. In patch-clamp experiments, the benzofuran KB 130015, shown in Fig. 16.11b, activated BK channels formed by hSlo (a) subunits in HEK 393 cells (Gessner et al., 2007). The channels were reversibly opened by shifting the open-probability/voltage relationship by about
60 mV. The shift was considerably stronger (
90 mV) with the addition of the Slo b1 subunit. Similar to mallotoxin, activation of BK channels was voltage dependent and calcium independent. In porcine pulmonary arteries contracted with prostaglandin F2a, KB 130015 induced endothelium-independent vasorelaxation, half-maximal at 43 mM.

FIGURE 16.11 The BK channel openers mallotoxin and KB 130013.

BK BLOCKERS/INHIBITORS

435

16.5 BK BLOCKERS/INHIBITORS 16.5.1

Anesthesia/Anesthetics

Deng and coworkers have studied the effects of the general anesthetic agent isoflurane, shown in Fig. 16.12 in oocytes expressing mslo1 (Wang et al., 2004). Using outside-out patches at different command potentials, it was determined that clinically relevant concentrations of isoflurane inhibited BK channels in a reversible, voltage-dependent manner. 16.5.2

Glaucoma

Ocular hypertension without optic nerve damage is believed to be a prelude to glaucoma, a condition in which sustained intraocular pressure results in optic nerve damage and irreversible loss of visual function. If elevated intraocular pressure is treated before nerve damage occurs, it may be possible to prevent the loss of visual function associated with glaucoma. BK channel blockers have been claimed in the patent literature to diminish fluid in the eye, thereby lowering intraocular pressure. Researchers at Merck have claimed a series of 70 paxilline (Fig. 16.13a) –like natural products as BK blockers for the treatment of glaucoma (Goetz et al., 2003). The compounds block BK channels in CHO cells constitutively expressing the a subunit, or in HEK 293 cells constitutively expressing both a and b1 subunits at compound concentrations at, or below, 100 nM. The inventors describe an in vivo model to measure intraocular pressure; however, no data on compound activity were provided. Another patent application claiming BK blockers for the treatment of glaucoma disclosed a series of indazoles, as illustrated in Fig. 16.13b (Doherty et al., 2004) The compounds blocked BK channels in transiently transfected TsA-201 cells as well as decreased BK currents in human nonpigmented ciliary epithelial cells. No biological data were presented that was specific to glaucoma. In a follow-up patent application to the indazoles, a series of tetrahydrocarbazoles, exemplified by Fig. 16.13c, were claimed as BK channel blockers, which lower intraocular pressure and therefore would be of use for the treatment of glaucoma (Gao and Shen, 2006). No other biological data were presented in this application.

FIGURE 16.12 The BK channel blocker isoflurane.

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FIGURE 16.13 BK channel blockers to treat glaucoma.

16.5.3

Malignant Glioma

BK channel blockers may have utility in the treatment of glioma (primary brain tumors believed to have originated from normal glial cells) (see Chapter 4). Gliomas account for 20% of all brain malignancies (Weaver et al., 2004), and are generally very difficult to treat with poor prognosis. In at least 50% of human gliomas, the epidermal growth factor receptors (erbB) are upregulated. The erbB receptors are tyrosine kinases that activate downstream signaling, resulting in an increase in intracellular calcium concentrations and activation of BK channels. The change in potassium conductance that results from the calcium-induced opening of BK channels has been shown to accompany the proliferation of numerous cell types. Examination of malignant glioma biopsy tissue revealed a significant expression of BK channel protein, which was subsequently identified as a novel splice variant of the hSlo gene with enhanced calcium sensitivity and named gBK (Liu et al., 2002; Ransom et al., 2002). Modulation of glioma BK channels was demonstrated by disruption of constitutive erbB2 activation. Application of Tyrphostin AG825, as shown in Fig. 16.14, a specific erbB2 inhibitor, induced a 30 mV positive shift in gBK channel activation in a dose-dependent manner in cell-attached patches (Olsen et al., 2005). The modulation appeared to occur via changes in intracellular calcium levels. A similar decrease in BK channel activation produced by a specific small-molecule blocker of these channels could therefore have utility in preventing glioma proliferation.

FIGURE 16.14 The glioma BK channel modulator tyrphostin AG 825.

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16.6 CONCLUSION BK channel modulation remains an elusive target for specifically developing drugs to treat human diseases. Although many compounds have demonstrated efficacy in animal models of diseases thought to be regulated by BK channels, to date none have reachedthemarket.However,thatisnottosaythattherearenoBKchannelmodulatorson the market. Drugs such as zonisamide and cilostazol were approved based on their demonstrated clinical efficacy in the treatment epilepsy and intermittent claudation, respectively. After their approval, further evaluation revealed the BK-modulating properties of both drugs, which may play a role in their mechanism of action. With the increasing knowledge of BK channel regulation, it should only be a matter of time before a drug specifically targeting BK channels reaches the market for one of the indications discussed in this chapter, or another yet uncharacterized utility.

REFERENCES Biagi G, Calderone V, Giorgi I, Livi O, Martinotti E, Martelli A, Nardi A, 2004a. 1,5Diarylsubstituted 1,2,3-triazoles as potassium channel activators. VI. Il Farmaco 59(5): 397–404. Biagi G, Giorgi I, Livi O, Nardi A, Calderone V, Martelli A, Martinotti E, LeRoy Salerni O, 2004b. Synthesis and biological activity of novel substituted benzanilides as potassium channel activators. V. Eur J Med Chem 39(6): 491–498. Boy KM, Guernon JM, Sit S-Y, Xie K, Hewawasam P, Boissard CG, Dworetzky SI, Natale J, Gribkoff VK, Lodge N, Starrett JJE, 2004. 3-Thio-quinolinone maxi-K openers for the treatment of erectile dysfunction. Bioorg Med Chem Lett 14(20): 5089–5093. Calderone V, 2002. Large-conductance, Ca2þ-activated Kþ channels: function, pharmacology and drugs. Curr Med Chem 9: 1385–1395. Calderone V, Giorgi I, Livi O, Martinotti E, Mantuano E, Martelli A, Nardi A, 2005. Benzoyl and/or benzyl substituted 1,2,3-triazoles as potassium channel activators. VIII. Eur J Med Chem 40(6): 521–528. Calderone V, Coi A, Fiamingo FL, Giorgi I, Leonardi M, Livi O, Martelli A, Martinotti E, 2006a. Structural modifications of benzanilide derivatives, effective potassium channel openers. X. Eur J Med Chem 41(12): 1421–1429. Calderone V, Fiamingo FL, Giorgi I, Leonardi M, Livi O, Martelli A, Martinotti E, 2006b. Heterocyclic analogs of benzanilide derivatives as potassium channel activators. IX. Eur J Med Chem 41(6): 761–767. Carl A, Bowen S, Gelband CH, Sanders KM, Hume JR, 1992. Cromakalim and lemakalim activate Ca2þ-dependent Kþ channels in canine colon. Pflugers Arch 421: 67–76. Cheney JA, Weisser JD, Bareyre FM, Laurer HL, Saatman KE, Raghupathi R, Gribkoff VK, Starrettt JE Jr, McIntosh TK, 2001. The maxi-K channel opener BMS-204352 attenuated regional cerebral edema and neurologic motor impairment after experimental brain injury. J Cereb Blood Flow Metab 21(4): 396–403. Clerc N, Furness JB, 2002. Intrinsic primary afferent neurones of the digestive tract. Neurogastroenterol Motil 16(Suppl 1): 24–27.

438

SMALL-MOLECULE MODULATORS

Doherty JB, Chen MH, Liu L, Natarajan S, Shen D-M, Tynebor RM, 2004. Ophthalmic compositions for treating ocular hypertension. Wo 2004/043354. Findlay I, Dunne MJ, Peterson OH, 1985. High-conductance Kþ channel in pancreatic islet cells can be activated and inactivated by internal calcium. J Membr Biol 83: 168–175. Gao Y-D, Shen D-M, 2006 Ophthalmic compositions for treating ocular hypertension. Wo 2006/044232. Gessner G, Heller R, Hoshi T, Heinemann SH, 2007. The amiodarone derivative 2-methyl-3(3,5-diiodo-4-carboxymethoxybenzyl)benzofuran (KB130015) opens large-conductance Ca2þ-activated Kþ channels and relaxes vascular smooth muscle. Eur J Pharmacol 555 (2–3): 185–193. Ghaemi SN, Zablotsky B, Flikowski MM, Dunn RT, Pardo TB, Isenstein E, Baldassano CF, 2006. An open prospective study of zonisamide in acute bipolar depression. J Clin Psychopharmacol 26(4): 385–388. Goetz MA, Kaczorowski GJ, Monaghan RL, Strohl WR, Tkacz JS, Merck & Co., Inc., USA, assignee. 2003 20030613. Maxi-K potassium channel blockers for treatment of glaucoma and as ocular neuroprotective agents. Application: WO patent 2003-US19013 2003105868. Gribkoff VK, Starrett JE Jr, Dworetzky SI, 1997. The pharmacology and molecular biology of large-conductance calcium activated (BK) potassium channels. In: August JT, Anders MW, Murad F, Coyle JT (Eds),Academic Press, San Diego, pp. 319–348. Gribkoff VK, Starrett JE, Dworetzky SI, Hewawasam P, Boissard CG, Cook DA, Frantz SW, Heman K, Hibbard JR, Huston K, Johnson G, Krishnan BS, Kinney GG, Lombardo LA, Meanwell NA, Molinoff PB, Myers RA, Moon SL, Ortiz A, Pajor L, Pieschl RL, PostMunson DJ, Signor LJ, Srinivas N, Taber MT, Thalody G, Trojnacki JT, Wiener H, Yeleswaram K, Yeola SW, 2001. Targeting acute ischemic stroke with a calcium-sensitive opener of maxi-K potassium channels. Nat Med 7(4): 471–477. Hewawasam P, Fan W, Knipe J, Moon SL, Boissard CG, Gribkoff VK, Starrett JE, 2002a. The synthesis and structure–activity relationships of 4-aryl-3-aminoquinolin-2-ones: a new class of calcium-dependent, large conductance, potassium (maxi-K) channel openers targeted for post-stroke neuroprotection. Bioorg Med Chem Lett 12(13): 1779–1783. Hewawasam P, Gribkoff VK, Pendri Y, Dworetzky SI, Meanwell NA, Martinez E, Boissard CG, Post-Munson DJ, Trojnacki JT, Yeleswaram K, Pajor LM, Knipe J, Gao Q, Perrone R, Starrett JE, 2002b. The synthesis and characterization of BMS-204352 (MaxiPost(TM)) and related 3-fluorooxindoles as openers of maxi-K potassium channels. Bioorg Med Chem Lett 12(7): 1023–1026. Hewawasam P, Ding M, Chen N, King D, Knipe J, Pajor L, Ortiz A, Gribkoff VK, Starrett J, 2003a. Synthesis of water-soluble prodrugs of BMS-191011: a maxi-K channel opener targeted for post-stroke neuroprotection. Bioorg Med Chem Lett 13(10): 1695–1698. Hewawasam P, Fan W, Ding M, Flint K, Cook D, Goggins GD, Myers RA, Gribkoff VK, Boissard CG, Dworetzky SI, Starrett JE, Lodge NJ, 2003b. 4-Aryl-3-(hydroxyalkyl) quinolin-2-ones: novel maxi-K channel opening relaxants of corporal smooth muscle targeted for erectile dysfunction. J Med Chem 46(14): 2819–2822. Hewawasam P, Fan W, Cook DA, Newberry KS, Boissard CG, Gribkoff VK, Starrett J, Lodge NJ, 2004. 4-Aryl-3-(mercapto)quinolin-2-ones: novel maxi-K channel opening relaxants of corporal smooth muscle. Bioorg Med Chem Lett 14(17): 4479–4482. Holland M, Langton PD, Standen NB, Boyle JP, 1996. Effects of the BKca channel activator, NS1619, on rat cerebral artery smooth muscle. Br J Pharmacol 117: 119–129.

REFERENCES

439

Hong KW, Kim KY, Lee JH, Shin HK, Kwak YG, Kim S-O, Lim H, Yoo S-E, 2002. Neuroprotective effect of (2S,3S,4R)-N00 -cyano-N-(6-amino-3, 4-dihydro-3-hydroxy-2-methyl-2-dimethoxymethyl-2H-benzopyran-4-yl)-N0 -benzylguanidine (KR-31378), a benzopyran analog, against focal ischemic brain damage in rats. J Pharmacol Exp Ther 301(1): 210–216. Hong KW, Kim KY, Shin HK, Lee JH, Choi JM, Kwak Y-G, Kim CD, Lee WS, Rhim BY, 2003. Cilostazol prevents tumor necrosis factor-{alpha}-induced cell death by suppression of phosphatase and tensin homolog deleted from chromosome 10 phosphorylation and activation of Akt/cyclic AMP response element-binding protein phosphorylation. J Pharmacol Exp Ther 306(3): 1182–1190. Hong K, Lee J, Kima K, Park S, Lee W, 2006. Cilostazol: therapeutic potential against focal cerebral ischemis. Curr Pharm Des 12(5): 565–573. Huang C-W, Huang C-C, Wu S-N, 2007. Activation by zonisamide, a newer antiepileptic drug, of large-conductance calcium-activated potassium channel in differentiated hippocampal neuron-derived H19-7 cells. J Pharmacol Exp Ther 321(1): 98–106. Kim KY,Lee JH,Park JH,Yoo M-A,Kwak Y-G,Kim S-O,Yoo S-E,Hong KW, 2004. Anti-apoptotic action of (2S,3S,4R)-N00 -cyano-N-(6-amino-3, 4-dihydro-3-hydroxy-2-methyl-2-dimethoxymethyl-2H-benzopyran-4-yl)-N0 -benzylguanidine (KR-31378) by suppression of the phosphatase and tensin homolog deleted from chromosome 10 phosphorylation and increased phosphorylation of casein kinase2/Akt/cyclic AMP response element binding protein via maxi-K channel opening in neuronal cells. Eur J Pharmacol 497(3): 267–277. Kunze WAA, Clerc N, Furness JB, Gola M, 2000. The soma and neurites of primary afferent neurons in the guinea-pig intestine respond differentially to deformation. J Physiol 526(2): 375–385. Latorre R, Oberhauser A, Labarca P, Alvarez O, 1989. Varieties of calcium-activated potassium channels. Annu Rev Physiol 51: 385–389. Lee D, Marino JP, Zhao Y, Smithkline Beecham Corporation, USA, assignee. 2005 20040715. Preparation of indolyl substituted benzoic acids as large conductance calcium activated potassium channel openers for treating urinary tract disorders. Application: WO patent 2004-US22706; 2005009993. Lerche H, Fahlke C, Iaizzo PA, Lehmann Horn F, 1995. Characterization of the highconductance Ca2þ-activated Kþ channel in adult human skeletal muscle. Pflugers Arch 429: 738–747. Liu X, Chang Y, Reinhart PH, Sontheimer H, 2002. Cloning and characterization of glioma BK, a novel BK channel isoform highly expressed in human glioma cells. J Neurosci 22(5): 1840–1849. McCann JD, Welsh MJ, 1986. Calcium-activated potassium channels in canine airway smooth muscle. J Physiol (Lond) 37: 113–127. Meech RW, 1978. Calcium-dependent potassium activation in nervous tissues. Annu Rev Biophys Bioeng 7: 1–8. Melman A, Bar-Chama N, McCullough A, Davies K, Christ G, 2006. hMaxi-K gene transfer in males with erectile dysfunction: results of the first human trial. Hum Gene Ther 17(12): 1165–1176. Nardi A, Calderone V, Chericoni S, Morelli I, 2003. Natural modulators of large-conductance calcium-activated potassium channels. Planta Med (10): 885–892. Ohwada T, Nonomura T, Maki K, Sakamoto K, Ohya S, Muraki K, Imaizumi Y, 2003. Dehydroabietic acid derivatives as a novel scaffold for large-conductance calcium-activated Kþ channel openers. Bioorg Med Chem Lett 13(22): 3971–3974.

440

SMALL-MOLECULE MODULATORS

Olsen MLW, Weaver AK, Ritch PS, Sontheimer H, 2005. Modulation of glioma BK channels via erbB2. J Neurosci Res 81(2): 179–189. Pedarzani P, Kulik A, Muller M, Ballanyi K, Stocker M, 2000. Molecular determinants of Ca2þdependent Kþ channel function in rat dorsal vagal neurones. J Physiol 527(2): 283–290. Power EC, Ganellin CR, Benton DCH, 2006. Partial structures of ketoconazole as modulators of the large conductance calcium-activated potassium channel (BKca). Bioorg Med Chem Lett 16(4): 887–890. Ransom CB, Liu X, Sontheimer H, 2002. BK channels in human glioma cells have enhanced calcium sensitivity. Glia 38(4): 281–291. Romine JL, Martin SW, Gribkoff VK, Boissard CG, Dworetzky SI, Natale J, Li Y, Gao Q, Meanwell NA, Starrett JE, 2002. 4,5-Diphenyltriazol-3-ones: openers of large-conductance Ca2þ-activated potassium (maxi-K) channels. J Med Chem 45(14): 2942–2952. Romine JL, Martin SW, Meanwell NA, Gribkoff VK, Boissard CG, Dworetzky SI, Natale J, Moon S,Ortiz A,Yeleswaram S,Pajor L,Gao Q,Starrett JE, 2007. 3-[(5-Chloro-2-hydroxyphenyl)methyl]-5-[4-(trifluoromethyl)phenyl]-1,3,4-oxadiazol-2(3H)-one, BMS-191011: opener of large-conductance Ca2þ-activated potassium (maxi-K) channels, identification, solubility, and SAR. J Med Chem 50(3): 528–542. Sakamoto K, Nonomura T, Ohya S, Muraki K, Ohwada T, Imaizumi Y, 2006. Molecular mechanisms for large conductance Ca2þ-activated Kþ channel activation by a novel opener, 12,14-dichlorodehydroabietic acid. J Pharmacol Exp Ther 316(1): 144–153. Sargent CA,Grover GJ,Antonaccio MJ,McCullough JR, 1993. The cardioprotective, vasorelaxant and electrophysiological profile of the large conductance calcium-activated potassium channel opener NS-004. J Pharmacol Exp Ther 266: 1422–1429. Sivarao DV, Newberry K, Langdon S, Lee AV, Hewawasam P, Plym MJ, Signor L, Myers R, Lodge NJ, 2005. Effect of 4-(5-chloro-2-hydroxyphenyl)-3-(2-hydroxyethyl)-6-(trifluoromethyl)-quinolin-2(1H)-one (BMS-223131), a novel opener of large conductance Ca2þactivated Kþ (maxi-K) channels on normal and stress-aggravated colonic motility and visceral nociception. J Pharmacol Exp Ther 313(2): 840–847. Starrett JE Jr, Dworetzky SI, Gribkoff VK, 1996. Modulators of large-conductance calcium activated potassium (BK) channels as potential therapeutic targets. Curr Pharm Design 2: 413–428. Starrett JE, Gillman KW, Bocchino DM, Bien J, Boissard CG, Michael B, Calandra V, Chen CPH, Chen Y, Dangle C, Dischino D, Gao Q, Hansel S, Hewawasam P, Huang S, Kang S-H, Lentz K, Lopez OD, Pajor L, Pendri Y, Pilcher G, Rand P, Provencal DP, Rao V, Sarsfield E, Schmitz WD, Sit S-Y, Wang H, Wu D, Xie K, Zhang L, Gribkoff VK, 2004. Maxi-K openers as neuroprotective agents. Abstracts of Papers, 227th ACS National Meeting, Anaheim, CA, March 28–April 1, MEDI-167. Suarez-Kurtz G, Garcia ML, Kaczorowski GJ, 1991. Effects of charybdotoxin and iberiotoxin on the spontaneous motility and tonus of different guinea pig smooth muscle tissues. J Pharmacol Exp Ther 259: 439–443. Vrudhula VM, Dasgupta B, Qian-Cutrone J, Kozlowski ES, Boissard CG, Dworetzky SI, Wu D, Gao Q, Kimura R, Gribkoff VK, Starrett JE, 2007. Atropisomeric 3-(b-hydroxyethyl)-4arylquinolin-2-ones as maxi-K potassium channel openers. J Med Chem 50(5): 1050–1057. Wang Y-W,Xiong Y-C,Deng X-M, 2004. Effects of isoflurane and ethanol on large conductance Ca2þ-activated Kþ channels. J Medical Colleges of PLA 19(3): 181–182, 186.

REFERENCES

441

Wang Y-J,Liu Y-C,Chang H-D,Wu S-N, 2006. Diosgenin, a plant-derived sapogenin stimulates Ca2þ-activated Kþ current in human cortical HCN-1A neuronal cells. Planta Med (5): 430– 436. Weaver AK, Liu X, Sontheimer H, 2004. Role for calcium-activated potassium channels (BK) in growth control of human malignant glioma cells. J Neurosci Res 78(2): 224–234. Werner ME, Zvara P, Meredith AL, Aldrich RW, Nelson MT, 2005. Erectile dysfunction in mice lacking the large-conductance calcium-activated potassium (BK) channel. J Physiol (Lond) 567(2): 545–556. Zakharov SI, Morrow JP, Liu G, Yang L, Marx SO, 2005. Activation of the BK (SLO1) potassium channel by mallotoxin. J Biol Chem 280(35): 30882–30887. Zografos P, Li JH, Kau ST, 1992. Comparison of the in vitro effects of potassium channel modulators on detrusor and portal vein strips from guinea pigs. Pharmacology 45: 216–230.

17 HIGH-THROUGHPUT SCREENING TECHNOLOGIES IN ION CHANNEL DRUG DISCOVERY Edward B. Stevens, Andrew D. Whyment, and J. Mark Treherne Pfizer Global Research and Development, Sandwich Laboratories (IPC 351), Sandwich, Kent, CT13 9NJ, UK

17.1 INTRODUCTION Ion channels regulate many diverse physiological processes including the electrical impulses that underlie sensory and motor functions in the brain; the control of contractile activity of the heart, skeletal, vascular, and visceral smooth muscle; and nutrient uptake, hormone secretion, cell replication, and development. There is accumulating genetic evidence for the importance of ion channels in many disease pathologies, and so the modulation of their function may be of value in treating an increasing variety of diseases. Consequently, there is considerable further potential in future ion channel drug discovery to uncover novel therapies that address unmet medical need. Selecting the most appropriate ion channels for drug discovery research is based upon the potential therapeutic relevance and toxicological implications of modulating the function of that channel. Analysis of the molecular classification of drug targets found that all known orally bioavailable drugs only bind to approximately 500 distinct molecular targets (Drews, 2000). Only around 5% of these molecular targets, however, were classed as ion channels. This relatively low percentage (e.g., compared with enzymes) probably has resulted from a lack of suitable starting points for medicinal chemistry and difficulties

Structure, Function, and Modulation of Neuronal Voltage-Gated Ion Channels, Edited by Valentin K. Gribkoff and Leonard K. Kaczmarek Copyright
2009 John Wiley & Sons, Inc.

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in configuring relevant screens to identify novel chemical series. The uses of existing ion channel drugs span many therapeutic areas, and this observation suggests that there is considerable further potential for ion channel drug discovery with new and existing targets. As the selective permeability and gating of ion channels are responsible for maintaining cell membrane potential and controlling cellular excitability (Armstrong and Hille, 1998), which is a key role in the functioning of all cells, it is not surprising that ion channels are such important drug targets (Ashcroft and Roper, 1993). It is estimated that approximately 300 pore-forming ion channel subunits are encoded in the human genome, giving rise to many more functional channels by combination with various auxiliary subunits. Potassium channels, for example, represent the largest and most diverse subgroup of ion channels and play a central role in regulating the membrane potential of cells (Kaczorowski and Garcia, 1999; Coghlan et al., 2001). Consequently, potassium channel inhibitors and openers offer significant therapeutic opportunities in cardiac, smooth muscle, neuronal, immune, and secretory systems (Liu et al., 1998). Where there has been an existing starting point for medicinal chemistry, ion channels have proved to be a highly tractable class of drug targets for the pharmaceutical industry (Treherne, 2002). Nevertheless, ion channels remain a relatively underexploited class of protein targets, mainly because of the absence of adequate screening technologies that enable pharmaceutical and biotechnology companies to evaluate focused libraries of compounds against potential ion channels of interest and then support medicinal chemistry to optimize the active compounds. Consequently, the progress of ion channel drug discovery is currently being constrained by the ability to identify lead compounds that can provide tractable starting points for medicinal chemistry. Recent advances in laboratory automation have brought significant opportunities to increase screening throughput for ion channel assays, but careful assay configuration to model drug–target interactions in a physiologically relevant manner is an essential consideration when selecting an assay for screening. Spectrums of ion channel screening platforms are described in this chapter, and some of these new technologies are transforming ion channel screening. A particular emphasis is placed on the mechanistic basis of drug interaction with the channel. For example, screening techniques should be able to detect compounds that interact with the channel by a preferred mode of action, so that novel “state-dependent” blockers, for example, can be discovered. This requires the assay system to generate relevant data from the compounds screened at the earliest stage of the drug discovery process. Ion channels should be fully integrated throughout the drug discovery process and not just introduced at the end of discovery to work out the mechanism of action of the drug and to identify unexpected toxicological implications. Advances in laboratory automation, instrumentation, and assay miniaturization have brought significant increases in the potential for screening throughput. For ion channel assays, however, careful assay configuration is more important than is generally required for, say, enzyme assays. A number of ion channel screening platforms are described in this review to provide some insights into the variety of formats available for high-throughput screening (HTS), together with some of their inherent advantages and limitations. Particular emphasis is placed on the mechanistic

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basis of drug–target interaction in the design of HTS approaches and how this relates to ion channel function. For example, an important aspect of the development of ion channels as drug targets is their ability to exist in multiple conformational states (e.g., open, closed, and inactivated) and show “state-dependent” drug sensitivity (Liu et al., 1998). This property of “state-dependent” blockade can be exploited therapeutically to discover drugs that selectively target the forms of the channel present under “pathological” conditions, while sparing normal channel functioning. The benefits of being able to select “state-dependent” ion channel modulators from screens can be of key importance in the discovery of novel medicinal chemistry and in avoiding potential toxicity complications. In this chapter, the focus is to update the reader on specific screening technologies and also attempt to illustrate how the technologies can be used to discover ion channel drugs.

17.2 ION CHANNEL HTS TECHNOLOGIES High-throughput, low false-positive hit rate, no false negatives, reliability, reproducibility, detection of fast channel activation and inactivation, good correlation with electrophysiology, low cost, and amenability to miniaturization are all characteristics that would make the ideal ion channel assay platform. Although there have been some very exciting developments over the last decade, still no single high-throughput ion channel assay platform is available that meets all these rigorous criteria. Current ion channel assay platforms can be broadly separated into several categories: ligand binding, ionic flux, fluorescence, luminescence, single-cell electrophysiology, and intact tissue electrophysiology. These different areas will be reviewed in the next sections. 17.2.1

Ligand Binding Assays

Many ligands have been identified that bind to ion channels or channel accessory proteins to activate or modulate their function. Typically, the ligand is radiolabeled and compounds are identified that competitively displaced or allosterically effected ligand binding to the ion channel. This approach usually identifies only compounds that behave in a mechanistically similar way to the labeled ligand. However, such techniques have been successfully used to discover calcium channel blockers and NMDA receptor antagonists. Ligand binding assays have also been used for discovering and validating some of the first-generation (e.g., tolbutamide, chlorpropamide, acetohexamide, and tolazamide) and second-generation (e.g., glibenclamide, glipizide, and glibornuride) sulphonylureas. Sulphonylureas stimulate insulin secretion by binding to sulphonylurea receptors leading to the closure of the ATP-sensitive potassium channel. Consequently, many sulphonylureas that were used in the treatment of maturity-onset diabetes were discovered by using binding assays. However, the ligand binding approach has limited success in discovering novel chemical series, as compounds that modulate ion channel activity independently to sulphonylurea binding go undetected. More recently, new improved classes of compound, unrelated

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to sulphonylureas, have been discovered using functional assays rather than the traditional ligand binding assays (Emilien et al., 1999). Ligand binding assays have also been used for identifying compounds with undesirable pharmacological properties. For example, compounds displaying toxicity toward the human ether-a-go-go-related gene (hERG) potassium channel may prolong the QT phase of a typical cardiac electrocardiogram (De Ponti et al., 2000). Long QT can result in the onset of ventricular arrhythmia and occasional sudden death. Dofetilide is a drug used to treat cardiac arrhythmias that functions by blocking hERG. Due to its hERG blocking capabilities, radiolabeled dofetilide can be used to identify other compounds that modulate hERG activity by performing radioligand binding assays, which has been patented. Radiolabeled dofetilide is incubated with whole cell or membrane preparations containing hERG, and compounds are then added and assessed for their ability to displace bound dofetilide (Diaz et al., 2004). However, compounds that modulate hERG activity via an alternative mechanism to dofetilide are not detected in this type of assay. This is a serious disadvantage considering the structural diversity of known compounds that modulate hERG activity (Vandenberg et al., 2001). Accordingly, functional electrophysiological assays for hERG safety pharmacology profiling are preferred when configuring a useful screen. Throughput of binding assays has been increased by moving away from filter binding assays toward homogenous assays that are more amenable to automation, for example, scillintation proximity assays (Zhang et al., 2006). The lower sensitivity of homogenous HTS assays has resulted in the development of radioligands with higher specific activity than tritiated radioligands, for example, [35S]MK-0499 for hERG binding (Raab et al., 2006). Fluorescence polarization (FP) is a technique based on the reduced tumbling rate of a fluorescent compound upon binding to the protein target, resulting in a polarized emission signal. FP binding assays have been adapted to GPCR HTS, providing a low cost, homogenous nonradiometric assay, especially in combination with “redshifted” fluorescent probes that reduce compound interference and “false-postive” rates (Allen et al., 2000). This technology is slowly being adapted to ion channel binding assay; for example, a hERG FP binding assay has been developed using a Cy3B labeled dofetilide derivative that shows a high correlation with data from [3H]-dofetilide binding (Deacon et al., 2007). However, further implementation of ion channel FP binding assays is limited by availability and development costs of highaffinity fluorescent probes. In summary, the application of ligand binding assays to ion channel pharmacology, although useful in certain specific examples, is relatively unattractive because of the limited information content and the inability to identify compounds that modulate ion channel activity independently by the mechanism of the labeled ligand. Further disadvantages include difficulties in modifying or synthesizing the ligand of interest and the influence of numerous parameters known to affect ligand binding. These include state-dependent ligand binding, voltage-dependent ligand binding and the binding of the ligand to multiple binding sites. In addition, some ion channels are not amenable to ligand binding assays, as there may be no known selective high-affinity ligands available. Applications outside of ion channel pharmacology and traditional screening include ion channel localization studies, isolation and identification of a

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ligand binding partner, and drug discovery platforms for the identification of compounds that may up- or downmodulate ion channel expression in the cell membrane (Ford et al., 2002). Nevertheless, where applicable, ligand binding assays are compatible with very high-throughput screening rates for ion channels (such as 100,000 data points in 24 h) at comparatively low cost per data point. 17.2.2

Ion Flux Assays

Ion channels are proteins that span the lipid bilayer of the cell membrane and provide an aqueous pathway through which specific ions such as sodium, potassium, calcium and chloride can pass (Choe and Robinson, 1998). The ability of channels to distinguish between different ionic species is a fundamental aspect of their function and is achieved by specific amino acid residues within the pore of the channel (Catterall, 1994; Sather et al., 1994; Bargmann, 1998). Flux assays exploit the fact that the pores of cation channels are permeable to other monovalent cations (Ford et al., 2002; Gill et al., 2003; Parihar et al., 2003), for example, the pore of a potassium channel is permeable to rubidium. By looking at the cellular efflux of potassium channel permeable ions, which are radioisotopic, one can monitor potassium channel activity. For potassium channels, radioactive rubidium 86Rbþ is the preferred isotope due to its high-energy emission characteristics that allows its quantification by Cerenkov counting without the requirement of liquid scintillation fluid addition. This radiometric flux assay principle has also been used to look at other ion channel types: 14C-guanidinium or 22Naþ for sodium channels, 45 Ca2þ for calcium channels, and 125I for chloride channels. Flux assays require a steady-state level of channel activity to carry significant ion flow to provide a robust assay. If the ion channel of interest is not active at resting membrane potential, the channel can be opened by various biochemical techniques. For example, calcium-activated channels can be activated by the addition of calcium ionophores, while voltage-gated channels can be activated using gating modifiers or elevated external Kþ concentrations (typically, 50 mM Kþ) to depolarize the cell. However, modifying the gating mode of the channel can have dramatic effects on pharmacology related to state-dependent interaction of the compound. For example, a high level of external Kþ reduces the affinity of dofetilde and cisapride binding to hERG, possibly by destabilizing C-type inactivation. Furthermore, the permeant tracer itself can affect channel function; for example, Rbþ reduces affinity of compounds for hERG by interacting with inactivation mechanisms (Rezazadeh et al., 2004). Safety and disposal concerns for high-energy radioisotopes are major limitations to configuring a screen for a large number of compounds. Atomic adsorption spectroscopy is an alternative nonradiometric assay system that is more amenable to automation and high-throughput screening. For example, radioactive 86 Rbþ can be substituted with nonradioactive 85 Rbþ and the amount of 85 Rbþ present in a given solution accurately quantified by atomic adsorption spectroscopy (Terstappen, 1999). The correlation between potency determinations between radiometric and these nonradiometric assays are virtually identical. Furthermore, atomic adsorption spectroscopy gives better sensitivity (increased signal-to-background ratio) compared

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to conventional radiometric detection methods. Although the majority of highthroughput nonradiometric ion flux screens have been developed for potassium channels using Rbþ efflux (Parihar et al., 2003; Wang et al., 2004; Gill et al., 2007), the technology is amenable to other cation channels such as sodium channels using lithium as the permeant tracer ion (see Figs. 17.1–17.3) and could be applied to calcium channels using strontium as the permeant ion. An indirect method of measuring Cl
channel activity has been reported using measurement of Agþ in solution after precipitation of insoluble AgCl following Cl
efflux (Gill et al., 2006). The throughput of nonradiometric flux assays is limited by the rate of measurement using atomic spectrometry. As current technology can measure only up to a dozen wells at once, miniaturization from 96 to 394-well format compromises assay quality without providing any advantage in throughput. A high-throughput reader for ion channel flux assays, ICR12000, has been designed by Aurora Biomed. The system comprises 12 atomic absorption spectrometers in parallel with a built-in plate stacker

FIGURE 17.1 Principles of Liþ influx assay for measuring Naþ channel activity using atomic emission spectrometry. (a) Cooperative effects of veratridine and scorpion venom on Nav1.5 channel kinetics measured using whole patch-clamp technique. Nav1.5 currents recorded in control conditions, 100 mM veratridine, 100 mM veratridine, and 5 mg mL
1Leiurus quinquestriatus hebraeus (Lqh) scorpion venom. Whole-cell patch-clamp recordings were performed on a HEK293–Nav1.5 cell line using an EPC9 amplifier with pulse v8.5. Currents were evoked using 10 mV increments from
100 to 40 mV for 50 ms from a holding potential of
100 mV. External solution contained 150 mM NaCl, 10 mM KCl, 3 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, and pH 7.4. Internal patch solution contained 160 mM KCl, 1 mM EGTA, 0.5 mM MgCl2, 10 mM HEPES, and pH 7.4. (b) Diagram illustrating basis of the Liþ influx assay. HEK293–Nav1.5 cells are incubated with or without compound for 10 min and then stimulated with 200 mM veratridine and 10 mg
1 mL Lqh scorpion venom for 60 min. Incubation buffer contains 139 mM LiCl, 10 mM HEPES, 1 mM MgCl2, 2 mM CaCl2, 5 mM glucose, and pH 7.4. Following washout of Liþ and Naþ channel activators, the cells are permeabilized using 0.1% SDS and the cell contents measured using the atomic emission mode of a ThermoElemental Solaar M series spectrometer connected to Gilson 222XL autosampler.

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FIGURE 17.2 Development of an Nav1.5 Liþ influx assay using atomic absorption spectrometry. (a) Veratridine/scorpion venom-induced Liþ influx is absent in HEK wild-type cells in comparison to Nav1.5 stably expressed in HEK cells (mean þ/
SD, n ¼ 12). (b) Liþ counts for positive (30 mM prenylamine) and negative (0.1% DMSO) control wells for a 45-plate screening run. (c) Z0 values calculated for each screening plate. (d) Correlation between duplicate IC50 values for compounds identified as hits from the LOPAC library. S. Liness, A. Cook and E. Stevens unpublished data.

and liquid handling system and claims measurements of up to 60,000 data points per day. An alternative method to read high numbers of samples is to integrate conventional atomic absorption spectometers with a robot arm, plate hotel, and liquid handling systems, for example, BioFocus DPI’s reader system integrated by Process Analysis and Automation Ltd (see ion channel studies by atomic absorption application note, www.paa.co.uk). The reader system can be further simplified by measuring atomic emission rather than atomic absorption (which is a more sensitive technique for measuring alkali metals); this removes the need for hollow cathode lamps. Atomic emission spectrometers have the added advantage of multielement analysis allowing measurement of ion flux from multiple ion channel types in the same screening plate. 17.2.3

Fluorescent Dye Assays

For ion channels, fluorescent dyes function in one of the two ways. First, dyes may be directly affected by the ion of interest, for example, calcium, halide, and thallium dyes. Second, dyes may indirectly monitor ion channel activity. For example, dyes that are

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FIGURE 17.3 Comparison of concentration–response data for prenylamine, quinidine, and lidocaine using whole-cell patch-clamp and Liþ influx assay. (a–c) Concentration–response data for prenylamine, quinidine, and lidocaine using whole-cell patch-clamp recordings of HEK293–Nav1.5 cells (mean þ/
SEM, n ¼ 3). Currents were evoked using a 10 ms pulse to
40 mV from a holding potential of
100 mV at 1 Hz. (d–f) Concentration–response data for prenylamine, quinidine, and lidocaine using Liþ flux of HEK293–Nav1.5 cells with atomic emission spectrometry (mean þ/
SEM, n ¼ 8). A. Cook, and E. Stevens unpublished data.

sensitive to changes in the cell resting potential can be used to assay the function of channels that control the membrane potential of a cell. Measurement of intracellular calcium concentration has mainly been developed to support screening of GPCRs (Monteith and Bird, 2005) but is amenable to Ca2þ channel and TRP channel screening (Witte et al., 2002). Single-wavelength dye measuring fluorescence intensity has been developed, for example, Fluo-4 (Invitrogen) or no-wash kits (comprising calcium dye with fluorescence quenchers) that increase throughput, for example, FLIPR Ca2þ assay kits (Molecular Devices). Single-wavelength dyes suffer from uneven dye loading, photobleaching, and dye leaking, which can be minimized using dualwavelength ratiometric dyes such as Fura-2 (Monteith and Bird, 2005). However, the majority of HTS labs use single-wavelength dye no-wash kits for simplicity and to increase throughput (Comely, 2006). Chloride channel assays have been configured by measuring halide quenching of quinolinium compounds (e.g., 6-methoxy-Nethylquinolinium iodide, MEQ), and pyrido[2,1-h]-pteridin-related compounds or the more successful (but patented) halide-sensing green fluorescent proteins (Galietta et al., 2001). On the contrary, Kþ channel assays have been developed by measuring thallium influx using the thallium-sensitive fluorescent dye benzothiazole coumarin

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acetoxymethyl ester (BTC-AM). This technology has been developed further by Molecular Devices to generate a no-wash dye kit using a novel dye ThalKal. Dyes that are sensitive to changes in the cell resting potential can be used to assay the function of channels that control the membrane potential of a cell. However, these conventional single-dye systems are susceptible to variation in dye loading, cell density, and cell volume. The use of a second dye during the assay allows ratiometric analysis and reduces artefacts, such as dye loading and cell density variation. Fluorescent cell-based assays that monitor ion channel activity often rely on membrane potential-sensitive dyes such as dis-(1, 2-dibutylbarbituric acid) trimethine oxonol (Di-BAC4(3)). Membrane potential-sensitive dyes are charged molecules that migrate across a lipid bilayer in a manner depending on the cell membrane potential. For example, a stable cell line expressing a voltage-gated potassium channel will be hyperpolarized and the negatively charged bis-oxonol dye DiBAC4(3) will be unable to migrate across the cell membrane into the cell. Blocking the voltage-gated potassium channel will set the resting membrane potential to a less hyperpolarized state, allowing DiBAC4(3) to migrate into the cell. Fluorescence produced from dye present in the cell can then be quantified by bottom plate reading on a conventional fluorescent plate reader in the presence of a cell impermeable dye quencher. Open potassium channels give low fluorescent counts, and blocked potassium channels give high fluorescent counts. Improved homogenous (by removing the wash steps) membrane-potential-sensitive dye systems developed by Molecular Devices using proprietary fluorescence masking or quenching compounds (Baxter et al., 2002) are faster, responding in less than 2 min, and are much less sensitive to temperature changes than DiBAC4(3). Fluorescence resonance energy transfer (FRET)-based dyes (commercially available from Invitrogen) are also used for looking at changes in membrane potential (Gonzalez et al., 2005). Briefly, an external voltage-insensitive donor dye (which is a coumarinlinked phospholipid) is FRET coupled to fast voltage-sensitive acceptor dye, such as DiSBAC2(3). When cells are in a hyperpolarized state, DiSBAC2(3) migrates out of the cell and FRET couples with the coumarin-linked phospholipid. For example, blocking potassium channels that control resting membrane potential causes the cells to become less hyperpolarized. DiSBAC2(3) migrates back into the cell, so that FRET (between donor and acceptor dyes) is unable to occur across the cell membrane. These FRETcoupled dye systems offer subsecond responses, ratiometric analysis, and reduced background fluorescence (since measurements are confined to the cell membrane). New brighter FRET donor dyes have been developed that unlike coumarin dyes are pH insensitive and have been used for screening ASIC channels (Maher and Wu, 2007). Modified cell-based fluorescent plate readers such as Molecular Device’s fluorometric imaging plate reader, FLIPRTETRA, Hamamatsu’s FDSS 6000 and PerkinElmer’s CellLux have been specifically developed for ion channel drug discovery including dual excitation for ratiometric measurements. These systems use a chargecoupled device (CCD) imaging system for simultaneous optical measurement on all wells of a 384- or 96-well microtiter plates and a simultaneous dispensing device for compound addition. High-energy dye excitation is from a water-cooled argon ion laser or xenon lamp. The FDSS 6000 system has been used for multiplexed assays where

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both membrane potential and Ca2þ mobilization are monitored in parallel, which could generate information-rich data for Ca2þ channel and TRP screening. Newer generation plate readers have now introduced built-in field stimulation capabilities for activating ion channels, which avoids having to simulate the channels using biochemical techniques, such as increasing extracellular potassium (Bugianesi et al., 2006; Huang et al., 2006). For example, FRET dye measurements of compounds acting at Nav1.3 using the modified VIPR system with integrated field stimulation, e-VIPR (Vertex Pharmaceuticals) shows a closer correlation with patch-clamp data than a VIPR FRET assay configured using sodium channel openers. In particular, FRET dye measurements of Nav1.3 using e-VIPR have been tested at different electrical stimulation frequencies and can identify use-dependent blockers such as lidocaine (Huang et al., 2006). 17.2.4

Flash Luminescence

Following the development of suitable readers and integrated HTS platforms, flash luminescence has been adopted for measuring Ca2þ mobilization assays for screening GPCRs (Comely, 2006). However, the technology can be extended to voltage-gated Ca2þ channels and ligand-gated channels (Dupriez et al., 2002; Di, 2007). The technique uses a bioluminescent Ca2þ-sensitive photoprotein such as aequorin as a reporter protein that is cotransfected with the target ion channel into a host cell line. Apoaequorin is the apoenzyme that is converted to the active enzyme aequorin in presence of its prosthetic group coelenterazine. Following Ca2þ binding, aeqourin oxidizes coelenterazine to coelenteramide, which results in emission of light measured as luminescence (Dupriez et al., 2002). The technique offers certain advantages over fluorescent assays such as lower cost, higher assay performance (low background and high signal-to-background ratio), requirement for lower cell numbers, and application to cells in suspension. For most ion channel assays, a cytoplasmic phosphoprotein is sufficient for detecting Ca2þ flux; however, rapidly inactivating Ca2þ channels or transient ligand-gated responses have Ca2þ flux kinetics that are too rapid to be detected by available instrumentation. Therefore, the commercially available photoprotein MitoPhotina (which has 70% similarity to aequorin) can be selectively targeted to mitochondria, which delays the detection of calcium waves by the photoprotein and slows the recording kinetics (Bovolenta et al., 2002). The MitoPhotina technology has been applied to P2X1 screening assays (Di, 2007). 17.2.5

Ion Channel Protein–Protein Interactions

Another approach to ion channel screening is to target compounds at sites of protein– protein interaction between or within subunits (Arkin and Wells, 2004), for example, reducing N-type inactivation by inhibiting docking of the inactivation ball or disturbing the interaction between a and b subunits. Wyeth has applied this approach to interaction of Kv4.2 with accessory subunit KChIP and interaction of Cav2.2 with Cavb3 subunits using a high-throughput yeast two-hybrid screen (Young et al., 1998; Bowlby et al., 2005). For example, the Cav2.2 screen used the Cavb3 coding sequence

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fused to the Gal4 DNA binding domain, while Cav2.2 intracellular loop domain I–II was fused to the Gal4 activation domain. Compound inhibition of binding of Cav2.2 to Cavb3 prevents reconstitution of Gal4 protein activity, preventing reporter gene expression (Young et al., 1998). A fluorescent protein–protein interaction screening assay has been developed by Leptus for Kv1.1-Kvb1 by immobilizing biotin-tagged Kvb1 core domains to streptavidin-coated plates and applying the T1 interaction domain of Kv1.1 labeled with Cy3 dye to the plates (Stafford et al., 2005). This technology has been extended to a screen for compounds that inhibit interaction of Cavb3 with Cav2.2 channels (Kanumilli et al., 2005). The emergence of label-free sensors, for example, Biacore (GE Healthcare) using surface plasmon resonance has provided another technique for monitoring affinity of protein–protein interactions and has been used to validate compounds identified from a yeast two-hybrid screen of compounds against Kv4.2–KChIP interactions (Bowlby et al., 2005). 17.2.6

Automated Single-Cell Electrophysiology

With the exception of FRET technology, the temporal response for the majority of the HTS assays is greater than a second. In contrast with these biochemical assays, electrophysiology is considered the “gold standard” for measuring modulatory effects of drugs on ion channels. Voltage-clamp recordings are direct measurements of channel activity and have a rapid temporal response (in milliseconds). Furthermore, voltage-clamp recordings are independent of the cell resting membrane potential, do not require modified external potassium concentrations to alter membrane potential and have exceptional signal-to-noise characteristics. Therefore, when using electrophysiology to study the action of compounds on ion channels, analysis can be performed under physiologically relevant timescales, voltages, and ionic conditions. Although the quality and reproducibility of electrophysiological data is substantially higher than data derived from biochemical assays, throughput by conventional electrophysiology is markedly lower. Therefore, considerable attention has been directed toward developing higher throughput electrophysiological assays and integrating these techniques earlier into the drug discovery process. However, the throughput of the systems that are currently available does not yet match the throughput of the biochemical assays that previously been described. One electrophysiological approach to increasing throughput has been to use two-electrode voltage clamp of the Xenopus oocyte heterologous expression system (Shih et al., 1998; Shieh et al., 2003). Some commercially available systems allow cDNA or cRNA injection and voltage-clamp recording from Xenopus oocytes in a 96-well plate, as well as a measuring head containing glass electrodes, reference electrodes, and perfusion ports. Some systems record from eight oocytes in parallel and have the potential of generating hundreds of data points per day and are suitable for higher throughput assays than are possible with conventional electrophysiology using mammalian cells. Such systems can be useful for quickly evaluating the function of a novel gene, but expression of mammalian ion channels in an amphibian expression system can often result in an abnormal pharmacology. Consequently, these systems are not often used for intensive screening or supporting medicinal chemistry.

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The electrophysiological technique used most commonly for more rigorous drug discovery is the whole-cell patch-clamp technique of mammalian cell lines transiently or stably expressing recombinant ion channels (Hamill et al., 2005). Under voltage-clamp conditions, measuring currents associated with the flow of charged ions monitors channel activity. The voltage-clamp technique is rapid and precise, allowing channel activity to be recorded over a millisecond timescale at different voltages. During the discovery process, electrophysiology has traditionally been used to validate HTS data and generate accurate IC50 or EC50 values later in lead optimization. Also, electrophysiology has been essential for determining mechanism of action of channel blockers by examining voltage dependence, state dependence, and frequency dependence. However, recently electrophysiology has been considered a potential platform for HTS by integrating it earlier into the drug discovery process where it can be used to explore hits discovered by other screening methods. Several automated electrophysiology systems have used a modification of conventional whole-cell patch-clamp technique where the entire patch-clamp system is miniaturized by removing the need for cell visualization, for example, Xention’s in-house “interface patching” (Treherne, 2002) and Wyeth’s RoboPatch (Vasilyev et al. 2006). Wyeth’s technology relies on blind patching of sedimented cells. In contrast, interface patching uses an inverted configuration where cells are loaded into a glass capillary and allowed to sediment to the air–solution interface, while a patch pipette is positioned beneath the capillary. A commercially available electrode-based automated electrophysiology system is Flyion’s “flip-the-tip” technology in which cells are placed within the electrode and subsequently flushed toward the tip where they form gigaseals (Lepple-Wienhues et al., 2003). The automated screening platform Flyion 8500 is capable of up to six recordings at once and has been adapted to recordings from intracellular organelles with diameters less than 1 mm, such as mitchondria. In contrast to conventional patch-clamp electrophysiology; however, these devices are higher throughput and largely controlled through software that controls the entire clamping process and allows the application of compounds and the analysis of electrophysiological data obtained from such drug applications. Chip-based planar patch-clamp systems dispense with the need for borosilicate patch pipettes (Bruggemann et al., 2003; Kutchinsky et al., 2003; Wang and Li, 2003; Wood et al., 2004) and instead use a substrate that has both optimal electrical insulation and membrane sealing behavior with machined apertures into which individual cells are positioned. Membrane currents are recorded across the aperture containing the cell using a current-to-voltage converter. Systems have been developed based on a planar patch-clamp system using, silicon, quartz, glass, polymer, or polydimethylsiloxilane (PDMS) chips (Sigworth and Klemic, 2005). There are currently a variety of commercially available systems for planar patch clamping. For example, Molecular Devices’ IonWorks currently provides the highest throughput using a 384-well recording chip (Schroeder et al., 2003). This level of throughput is achieved by compromising the quality and continuity of the patch-clamp recording. The system uses a perforated patch-clamp configuration with a low seal resistance (typically 50–100 MW) and no

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compensation for capacitance artefacts or series resistance compensation. During compound addition, the recording head is removed to allow pipettor access; therefore, voltage clamp is interrupted during recordings that results in cell depolarization and change in channel state, The system is designed for single-compound applications, therefore, data are analyzed in a similar way to conventional high-throughput screening using positive/negative controls and Z0 values to determine assay performance statistically. IonWorks HT compensates for recording failure rate by applying one compound to four recording sites; however, the system has recently been modified by using a “population patch-clamp” technique (IonWorks Quattro), with 64 apertures in each recording site resulting in a current recorded from multiple cells. This has a dramatic effect on success rate as the technique accommodates a low percentage of apertures unoccupied by cells. Therefore, only one recording site is required for each compound resulting in a fourfold greater throughput than IonWorks HT (Finkel et al., 2006). As each recording is constructing from multiple cells, stable cell lines with a high percentage of nonexpressors can be successfully screened. In addition, multiplexing with a mixture of different ion channel cell lines on the same recording site is possible (Dale et al., 2007). Planar patch-clamp systems based on high-quality recordings using both “gigaseal” formation and whole-cell recording configuration are available, for example, Sophion’s QPatch, Molecular Device’s PatchXpress, and Nanion’s Patchliner. These systems use uninterrupted recordings and are configured for sequential additions of compounds with intervening washout periods or addition of cumulative concentrations of compounds to single cells. PatchXpress uses Aviva Biosciences’ Sealpatch planar patch-clamp technology that consists of 50–100 mL wells with 1–2 mm diameter planar patch-clamp holes using PatchXpress wash probes (consisting of combined inlet and outlet channels) to washout compounds., where 90% solution replacement takes several hundred milliseconds (Xu et al., 2003; Tao et al., 2004). The newer generation of planar patch-clamp plates such as Sophion’s QPlate and Nanion’s NPC-chips uses integrated microfluidics that provide faster rates of solution exchange (